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f | Volume 48 LOGS (oe is Number 1 


7 ISSN 0024-0966 
a JOURNAL 


of the 


_ LEPIDOPTERISTS’ SOCIETY 


Published quarterly by THE LEPIDOPTERISTS’ SOCIETY 


Publié par LA SOCIETE DES LEPIDOPTERISTES 
Herausgegeben von DER GESELLSCHAFT DER LEPIDOPTEROLOGEN 
Publicado por LA SOCIEDAD DE LOS LEPIDOPTERISTAS 


Ma 3 23 February 1994 


THE LEPIDOPTERISTS’ SOCIETY 
EXECUTIVE COUNCIL 


PAUL A. OPLER, President JORGE E. LLORENTE-BOSQUETS, 
Ray E. STANFORD, Immediate Past Vice President 

President FREDERICK W. STEHR, 
CHEN-SHING LIN, Vice President Vice President 
JULIAN P. DONAHUE, Acting Secretary ROBERT J. BORTH, Treasurer 
Members at large: 
Charles V. Covell, Jr. Eric H. Metzler John V. Calhoun 
Linda S. Fink Robert K. Robbins Robert C. Lederhouse 
Scott E. Miller J. Benjamin Ziegler William E. Miller 


EDITORIAL BOARD 


PAUL A. OPLER (Chairman), FREDERICK W. STEHR (Member at large) 
JOHN W. BROWN (Journal), WILLIAM E. MILLER (Memoirs) 
STEPHANIE S. MCKOWN (News) 


HONORARY LIFE MEMBERS OF THE SOCIETY 


CHARLES L. REMINGTON (1966), E. G. MUNROE (1973), 
ZDRAVKO LORKOVIC (1980), IAN F. B. COMMON (1987), JOHN G. FRANCLEMONT (1988), 
LINCOLN P. BROWER (1990), DOUGLAS C. FERGUSON (1990), 
HON. MIRIAM ROTHSCHILD (1991), CLAUDE LEMAIRE (1992) 


The object of the Lepidopterists’ Society, which was formed in May 1947 and for- 
mally constituted in December 1950, is “to promote the science of lepidopterology in all 
its branches, .... to issue a periodical and other publications on Lepidoptera, to facilitate 
the exchange of specimens and ideas by both the professional worker and the amateur 
in the field; to secure cooperation in all measures” directed towards these aims. 

Membership in the Society is open to all persons interested in the study of Lepi- 
doptera. All members receive the Journal and the News of the Lepidopterists Society. 
Institutions may subscribe to the Journal but may not become members. Prospective 
members should send to the Treasurer full dues for the current year, together with their 
full name, address, and special lepidopterological interests. In alternate years a list of 
members of the Society is issued, with addresses and special interests. There are four 
numbers in each volume of the Journal, scheduled for February, May, August and 
November, and six numbers of the News each year. | 


Active members—annual dues $25.00 
Student members—annual dues $15.00 
Sustaining members—annual dues $35.00 
Life members—single sum $500.00 
Institutional subscriptions—annual $40.00 


Send remittances, payable to The Lepidopterists’ Society, to: Robert J. Borth, Treasurer, 
6926 North Belmont Lane, Fox Point, WI 53217, U.S.A.; and address changes to: Julian 
P. Donahue, Natural History Museum, 900 Exposition Blvd., Los Angeles, CA 90007- 
4057 U.S.A. For information about the Society, contact: Julian P. Donahue. To order 
back issues of the Journal, News, and Memoirs, write for availability and prices to the 
Publications Manager: Ronald Leuschner, 1900 John St., Manhattan Beach, CA 90266- 
2608, U.S.A. 


Journal of the Lepidopterists’ Society (ISSN 0024-0966) is published quarterly for 
$40.00 (institutional subscription) and $25.00 (active member rate) by the Lepidopterists’ 
Society, % Los Angeles County Museum of Natural History, 900 Exposition Blvd., Los 
Angeles, CA 90007-4057. Second-class postage paid at Los Angeles, California and ad- 
ditional mailing offices. POSTMASTER: Send address changes to the Lepidopterists’ 
Society, % Natural History Museum, 900 Exposition Blvd., Los Angeles, CA 90007-4057. 
If you have used a Society address many years after its publication date, and your mail 
is returned as undeliverable, please redirect it to the Natural History Museum address 
provided above. 


Cover illustration: The black swallowtail, Papilio polyxenes asterius (Stoll), is a wide- 
spread species of the eastern United States. The larvae feed on a variety of umbelliferous 
plants (Apiaceae). Original drawing by Erik Russell Wild, Museum of Natural History, 
University of Kansas, Lawrence, Kansas 66045-2454. 


JOURNAL OF 


Tue LEPIDOPTERISTS’ SOCIETY 


Volume 48 1994 Number 1 


Journal of the Lepidopterists’ Society 
48(1), 1994, 1-7 


PRESIDENTIAL ADDRESS 1993: ON THE COMPARATIVE 
DISTRIBUTIONS OF LEPIDOPTERA AND LEPIDOPTERISTS 


RAY E. STANFORD 
720 Fairfax Street, Denver, Colorado 80220, USA 


Both Lepidoptera and lepidopterists, each part of a much larger group 
of similar organisms, represent but fleeting moments in time. But the 
Lepidoptera have flitted a great deal longer than we have as their 
students, by four orders of magnitude. I have been interested in the 
spatial and temporal distribution of butterflies for nearly 50 years, 
beginning in the neighborhood of my house and expanding incremen- 
tally to all of western North America (Stanford & Opler 1993), but 
began to notice a consistent artifact on the dot maps: butterflies are 
shown to be more common in cities, along paved roads, and in beautiful 
mountain meadows than in deserts, on farms, or in abandoned mine- 
fields. This presentation is an attempt to put these biases into perspec- 
tive, based on several years’ attempts by me and others to find out what 
species do occur (and even thrive) in the less-well-known habitats of 
our large region. Before summarizing my own research in this realm, 
it is necessary to give some historical perspective both on bugs and 
buggers. 

Although some insect orders are known from fossils dating from early 
Cretaceous time, the first Lepidoptera appeared along with flowering 
plants about 120 million years before the present (mybp), and probably 
all families of Lepidoptera that exist today had differentiated by 66 
mybp (Emmel et al. 1992). Most fossil butterflies date from the late 
Eocene to early Oligocene epochs, about 48-34 mybp, or later. Exten- 
sive movements of species occurred during and following the Pleisto- 
cene glaciations, as has been determined both from examination of 
fossils and study of today’s distributions taking geologic evidence into 
consideration. Humans undertook equally extensive movements at the 
same time, extending into much of North and South America from 


2 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Eurasia perhaps as long ago as 35 thousand ybp. Lepidoptera appear 
in pictographs and caves dating from then, but it would be a stretch 
to refer to the artists as lepidopterists! The first drawings that are fairly 
easily determined to species are from Egyptian and Sumerian tombs 
from 5000 to 3000 ybp; Danaus chrysippus can be dated from drawings 
in Luxor created about 3500 ybp (Larsen 1990). The first surviving 
descriptions of Lepidoptera which may be considered scientific were 
by the Greek philosopher /scientist Aristotle, who lived and wrote in 
the 4th century BC. His accounts are sufficiently detailed that several 
species of butterflies and moths which still occur in his country could 
be considered described by him, but he failed to assign them Latin 
names or designate type localities! Common names (in Greek for chry- 
sippus) are entirely appropriate for common species (Miller 1992), but 
are a conundrum for uncommon ones (Scott 1993) for which every 
author seems to make up a different common name. For example, what 
should be ‘““Edwards’ skipper’? He described 51 species of them (as 
presently classified) from the western United States alone. I shall not 
attempt to review the history of lepidopterists since the time of Linnaeus 
(1753), Cramer (1775), or Fabricius (1807), but the late F. Martin Brown 
wrote many historical papers in addition to his monumental series on 
the W. H. Edwards taxa. Brown influenced many of us over several 
decades, and I find it appropriate to dedicate my remarks here to his 
memory since he was long a guiding force behind my research. 

Many factors influence the distribution of butterflies in space and 
time, including climate, host plants and other biological requirements, 
and the effects of human activity. In order for range maps to show the 
actual distribution of a species for any given interval of time, these 
factors and the potential biases and artifacts mentioned at the outset 
must be considered carefully. Also, the mapper must beware of intro- 
ducing errors by the very process of making maps (Monmonier 1991), 
and the changes in a species’ range over time require either several 
maps or different symbols denoting different time periods on a single 
map (e.g., Heath 1970). The British Atlas (Heath 1970) also shows 
different intensities of observation/collecting among the thousands of 
10 km grid squares, so that the presence or absence of a species in a 
certain area may be evaluated in terms of observation density as well 
as other factors. 

First off, I shall address the issue of errors which originate from the 
mapping process itself. All maps tell little white lies of necessity. Most 
of us grew up seeing Mercator-projection maps on classroom walls, 
where Greenland appears larger than the United States, and Antarctica 
is as long as the equator, but we learned quickly to adapt to these “lies.” 
My well-worn Colorado highway map shows a prominent north-south 


VOLUME 48, NUMBER 1 5 ° 


ribbon about 2 mm wide running from Cheyenne, Wyoming, to Raton, 
New Mexico, Interstate Highway 25. If that width were to scale, a DC- 
10 could easily land crosswise and never see or hit an automobile! Similar 
types of misinterpretation are possible on dot maps of small scale, where 
only a single dot in the center of each county (or other unit) will fit. 
For example, a common species such as Vanessa cardui, known from 
all counties of both Kansas and Nevada, appears to be very much more 
common in Kansas (105 tiny counties) than in Nevada (18 gigantic ones 
and 4 tiny ones) simply because the dots are nearly confluent in Kansas. 
Also, Boloria acrocnema shows in 4 large Colorado counties, with a 
total area of thousands of square miles, but the insect actually occurs 
in only a few several-acre colonies above treeline on the San Juan massif. 
Of course, in a scientific paper addressing either of these issues, the 
maps would be prepared in different formats, with equal size grid 
squares for the painted lady in Kansas and Nevada, and a large scale 
map to show the specific localities for the Uncompahgre fritillary in 
Colorado. In a work with over 1000 identical-format maps, these matters 
must be summarized by a simple caveat in the introduction: Beware 
of [implied] lies! Most range maps, whether showing discrete dots or 
shading, are based on county records in the United States because the 
county of the locality on the specimen label is usually easy to determine 
given the other label data and ready access to historical maps; also there 
is no standard grid system used among disciplines, although longitude/ 
latitude could be used. However, using counties as the basis requires 
considerable care to avoid plotting errors. Counties sometimes come or 
go, or change boundaries, or even names, and the names of towns 
correlate poorly with county names. Here are some examples. Before 
Colorado became a State in 1876, Denver was in Montana! That is, 
Montana Co., Kansas Territory. Several butterfly species were described 
from “Denver” which do not occur anywhere near the city and certainly 
didn’t then either, so a dot must not be placed there without better 
information. Grand Co., Utah Territory, included what are now Grand 
Co., Colorado, Grand Co., Utah, and nearly everything in between. In 
more recent years, Washabaugh Co., South Dakota, vanished into Jack- 
son Co., South Dakota (combining the map dots was quite easily ac- 
complished), while Yuma Co., Arizona, and Valencia Co., New Mexico 
split into 2 counties each (we had to go back to ground zero for many 
records), and Denver Co., Colorado gobbled up a lot of real estate in 
Adams Co. (requiring transfer of a few county dots). Bullfrog Co., 
Nevada, was created and then abolished so quickly that no action was 
necessary on our part. Several of the Sierra foothill counties in California 
changed their boundaries almost weekly in the days of the Gold Rush 
and afterward, but have fortunately been stable during most of col- 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


a 


LEPIDOPTERISTS, 1800 


CABBAGE WHITE (P. RAPAE), 1800 


M 
ster 


eB 54s 


LEPIDOPTERISTS, 1956 


CABBAGE WHITE (P. RAPAE), 1956 


(Published records) 


list) 


(Lepidopterists' Society membership 


1983-1993) 


LEPIDOPTERISTS, 1993 (Season Summaries 


1993 


CABBAGE WHITE (P. RAPAE), 


VOLUME 48, NUMBER 1 ) 


lecting efforts there since the time of Lorquin. The Sonora blue, de- 
scribed from “Sonora,” was from near the California gold camps, not 
Mexico! Then, beware of ambiguous and misleading names: Bent, Col- 
orado, is in Las Animas Co., while Las Animas is the seat of Bent Co.: 
also Cheyenne Co. is in Nebraska, while Cheyenne, Wyoming is in 
Laramie Co., and Laramie, Wyoming is in Albany Co. (NOT New 
York)! 

Secondly, maps reflect the habitats of lepidopterists at least as much 
as those of the Lepidoptera they study, and also the goals and biases 
of the students. I mentioned earlier that populated, easy-access, and 
beautiful places tend to be better known than unpopulated, remote, 
and barren ones, but another principle has been called Powell’s Law 
for its perpetrator J. A. Powell: distant places are more thoroughly 
studied than close ones, or “No field biologist does any significant work 
closer than 1000 miles from his home!’ A case in point could be the 
Chiricahua Mtns of Arizona, or the Galapagos Islands, but if one looks 
at the evolving knowledge of common species’ distributions, the exact 
opposite seems to be true. I have chosen the cabbage white, Pieris 
rapae, to illustrate this point. Panel A of Fig. 1 shows the range of this 
introduced Eurasian species in 1800, several years before it first ap- 
peared on our continent, side-by-side with the range of serious lepi- 
dopterists in the same year in the western United States: both zero! 
Panel B shows the known locations for each group in 1956, again nearly 
the same, in cities and towns only. Panel C shows where they are 
documented to exist in 1993—again identical, but this time nearly 
everywhere. So my corollary to Powell’s Law is the converse: “No one 
notices cabbage whites except in his/her own back yard!’ Except that 
a certain map-dotter finally picked one up on the west side of Loveland 
Pass, Colorado, 12,000’ above sea level, to complete the dots for all 63 
counties and prove Powell correct after all. The distribution of lepi- 
dopterists also is shown by the fact that 110 butterfly species are known 
from Scott Co., Kansas, compared with 30 to 45 in surrounding and 
equally depauperate counties, because Virgil Calkins lived and recorded 
species in his back yard in Scott City for several decades this century. 
In his honor I have therefore formulated Calkins’ Law: If one studies 
a habitat for long enough, more than 100 butterfly species will be found 
there. Anywhere! Another artifact is caused by the interests of the 
observers. Swallowtails and fritillaries are much better known (but 
probably NOT more widespread) than skippers or noctuid moths, be- 


—_— 


Fic. 1. County records for the cabbage white (Pieris rapae) (left column) and the 
distribution of lepidopterists (right column). A) 1800; B) 1956; C) 1993. 


6 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


cause they are more popular. Methods of observers affect the results 
also; like authors who accept literature records or sightings show more 
polka dots than those who accept only captures with voucher specimens 
and extremely detailed label data (frequently only their own). Iam not 
faulting careful work, and am sometimes annoyed at papers that show 
everything from soup to nuts thereby conferring a measure of credibility 
on frank errors that should be expunged, but I support a middle ground. 
Maps that are revised frequently, or are on computer files, can be 
updated to show deletions and corrections as well as additions and 
format changes. 

Finally, I pose a somewhat rhetorical question: When do range maps 
reach their maximal utility? An important point is reached when all 
common species are known from all or most grid squares or counties, 
because it can be inferred then that the ranges of rarer species are well- 
known also since most biologists are more interested in them, and that 
a scattergram stray species (such as Eurema mexicana) has not been 
encountered very frequently but can pop up nearly anywhere. We are 
nearing that inflection point for western butterflies now, and a map 
showing E. mexicana for all counties from Canada to Mexico in a few 
centuries would tell a different sort of lie, unless the relentless march 
of Eurasian weeds and fungi into the region—replacing most native 
plant species—should allow this particular species to flourish continent- 
wide. Will the range maps for cabbage whites in the year 2098 in 
western North American look very much like today’s, with a few more 
squares filled in, or will they look like those in 1800? In either case, 
the comparative distributions of common Lepidoptera and common 
people will finally be identical, but where will the uncommon ones be? 
Hopefully the uncommon lepidopterists will still be pursuing the un- 
common insects, as Rindge urged in his 1965 presidential address, which 
I had the privilege to hear in person. Collect NOW, he urged, because 
it soon would be too late. His message is still true, and even more urgent 
than nearly 30 years ago. Much collecting is possible with a good 
telephoto camera, but museums and universities still need material, 
especially of yet unknown or poorly known species, so the process of 
obtaining permits to collect specimens is well worth the effort. When 
all that is left is cabbage whites, a permit will probably not be required. 
If so, the butterflies may have to issue it. 


LITERATURE CITED 


EMMEL, T. C., M. C. MINNO & B. A. DRUMMOND. 1992. Florissant butterflies: A guide 
to the fossil and present-day species of central Colorado. Stanford University Press, 
California. x plus 118 pp., 9 color plates. 

HEATH, J. 1970. Provisional atlas of the insects of the British Isles, Part 1: Lepidoptera 
(Rhopalocera, Butterflies). Biological Records Centre, Monks Wood Experimental 


~I 


VOLUME 48, NUMBER 1 


Station, Abbots Ripton, Huntingdon, England, front matter plus index plus 57 full 
page maps. 

LARSEN, T. B. 1990. The butterflies of Egypt. American University in Cairo Press. 112 
pp., 8 color plates. 

MILLER, J. Y. (ed.). 1992. The common names of North American butterflies. Smith- 
sonian Institution Press, Washington, D.C. ix plus 177 pp. 

MONMONIER, M. 1991. How to lie with maps. University of Chicago Press. xi plus 
176 pp. 

RINDGE, F. H. 1965. Presidential Address 1965: The importance of collecting—Now. 
J. Lepid. Soc. 19(4):193-195. 

ScoTT, J. A. 1993. The common names of North American butterflies (book review). 
J. Lepid. Soc. 47(2):170-171. 

STANFORD, R. E. & P. A. OPLER. 1993. Atlas of Western U.S.A. butterflies, including 
adjacent parts of Canada and Mexico. Privately published. x plus 275 pp. 


Received and accepted for publication 20 October 1993. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 8-23 


A NEW GENUS OF WINTER MOTHS (GEOMETRIDAE) FROM 
EASTERN CALIFORNIA AND WESTERN NEVADA 


JERRY A. POWELL 
Essig Museum of Entomology, University of California, Berkeley, California 94720, USA 


AND 


DOUGLAS C. FERGUSON 


Systematic Entomology Laboratory, ARS, USDA, 
Washington, District of Columbia 20560, USA 


ABSTRACT. Tescalsia, a new genus of Geometridae, is described and assigned to 
the subfamily Larentiinae. It is represented by two new species: Tescalsia giulianiata 
Ferguson, known from 3 localities in and adjacent to the Owens Valley, Inyo Co., 
California, and T. minata Ferguson, from Mineral Co., Nevada. The female of T. giu- 
lianiata has linear, straplike forewings, vestigial hindwings, and long, slender legs that 
enable agile climbing in shrubs; the female of T. minata is unknown. Both sexes lack the 
proboscis and tympana. Adults of T. giulianiata are active at sundown and nocturnally 
in November and December, despite near freezing temperatures. 


Additional key words: brachypterous, flightless, Larentiinae, tympana. 


In 1976 Mr. Derham Giuliani, a keen naturalist of Big Pine, Cali- 
fornia, brought a brachypterous moth to Berkeley that he had collected 
in Deep Spring Valley, east of Big Pine, in early December 1978. So 
bizarre was the specimen, with peculiarly bristled, straplike forewings 
and vestigial hindwings and mouthparts, that Powell could not identify 
it to family. The taxonomic placement remained a mystery after Fer- 
guson and other lepidopterists at the National Museum of Natural 
History (NMNH) examined the specimen in 1977. 

Two additional females were taken in pitfall traps at Deep Spring 
Valley in December 1978, one of which laid eggs that produced first 
instar geometrid larvae. After additional visits by Giuliani, Powell, and 
others during November and December 1978-82, we had assembled 
9 winged males and 8 females, a sufficient sample to provide con- 
vincing circumstantial evidence for the association of the sexes. The 
males indicate that the mystery moth is an undescribed species best 
assigned to the Larentiinae (Geometridae). 

Later, males of a congeneric species, from Mina, Mineral Co., Ne- 
vada, that had been collected in 1914, were discovered by Ferguson in 
unidentified material at the NMNH. Although descriptions of the new 
genus and species were written several years ago, we delayed publi- 
cation, anticipating that the larva and host plant might be discovered; 
but that hope has not been realized. Hence, we present the descriptions, 
and characterize the egg, together with observations on the habitats 
and adult behavior. 


VOLUME 48, NUMBER 1 9 


MATERIALS AND METHODS 


Trapping method. The first female was observed on the sand, and 
a few males were netted near sundown; but most of the series was taken 
in pitfall traps. This technique is widely used for sampling ground- 
dwelling insects in various habitats and is particularly effective for 
nocturnal insects such as many beetles. Cups may be deployed empty, 
baited with truly disgusting materials, or partially filled with anti-freeze 
(ethylene glycol), which allows long-term sampling. If deployed empty, 
traps need to be checked frequently because many predators, spiders, 
scorpions, carabid beetles etc., are trapped. 

We used 9 oz. squat plastic tumblers (7 cm deep and 9 cm diameter 
at the rim); they are inexpensive and nest in compact packages for 
transport. We deployed them in transect lines of 50-100, about 2 m 
apart, usually situated at the bases of shrubs. We trapped with empty 
cups during single nights (when two of three females were killed by a 
predaceous mite and spider) or with anti-freeze over a several day 
period. Specimens taken from the latter were washed in water, then 
transferred to alcohol, and later dried for pinning. Understandably, 
lepidopterists rarely are familiar with the method, but it is an effective 
one for brachypterous forms. Winged males are also sometimes trapped, 
and most of our male Tescalsia were collected this way. The holotype 
was taken in early morning in an empty trap about 50 m from the 
nearest trapped female. 


SYSTEMATICS 
Tescalsia Ferguson, new genus 


Type species: Tescalsia giulianiata Ferguson, new species. 


Diagnosis. Characterized by the following combination of unusual 
characters: 1) in male, veins Sc and Rs fused for half length of hindwing, 
forking just before end of the very long cell, and beyond that point Rs 
and M, stalked together for “4 or % of distance from end of cell to outer 
margin; 2) female with greatly reduced forewing, linear and straplike, 
with numerous setae longer than width of wing along entire length of 
both fore and hind margins; 3) female with hindwing present only as 
a vestige concealed beneath base of forewing; 4) tympanic cavities in 
base of abdomen, characteristic of nearly all Geometridae, missing in 
both sexes; 5) proboscis missing in both sexes, as is true of some other 
winter moths. 

Description. MALE: Length of forewing: 12.0-17.5 mm. Head: An- 
tenna bipectinate, extremely delicate, with slender shaft and long, wide- 
ly spaced, setose branches and large scape 3-4 times thickness of shaft; 


10 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


shaft scaled dorsally. Labial palpus short, hardly extending beyond 
front. Eye not very large, its vertical length almost equal to width of 
front at narrowest point, but strongly protuberant in the type-species, 
less so in the other. Ocellus absent. Front protuberant, rounded. Chae- 
tosema normal, moderately to well developed, the two chaetosemata 
not extended transversely to meet behind head as in many Larentiinae. 
Tongue absent. Vestiture untufted. Legs slender, normal except that 
foretibia is extremely reduced and bears a large, conspicuous, double 
claw (Fig. 9), and foretarsus much more elongated than mid- or hind- 
tarsus; hindtibia with one or two pairs of spurs. 

Fore- and hindwing somewhat elongated, produced especially toward 
apex; tornus of both wings rounded; forewing length/width ratio about 
2.4/1.0, narrower than the 2.0/1.0 ratio of Operophtera (length = base 
to apex; width = length of line through anal angle meeting costa at 
90°); wing pattern mostly diffuse and indistinct, variable between and 
within species; hindwing maculation differing from that of forewing 
in its reduced pattern consisting of discal spot only or discal spot and 
diffuse postmedial band only; fringes of both wings unusually long; 
wings covered with wide, tulip-shaped to almost round scales, dentate 
with 3-5 short points distally or simple. Venation as described and 
figured (Fig. 8); Sc of forewing free from radials; discal cells long, that 
of forewing more than half and of hindwing about half length of wing, 
closed off almost straight transversely at ends; forewing with two large 
accessory cells; hindwing with discal cell unusually wide because of 
long fusion of Sc and Rs, with M, a fully developed, tubular vein, and 
with lst A a well-developed fold, 2nd A a well-developed vein, and 
3rd A entirely lost. Frenulum well developed and very long. 

Male genitalia (Figs. 10, 11). Valva and uncus simple; uncus ap- 
pearing to be fused to tegumen, forming a solid unit; gnathos absent; 
transtilla distinct, forming slender, complete bridge; manica spinulate, 
heavily so in one species, flanked by pair of knoblike, setose processes, 
the derivation of which is unclear; each knoblike process with delicate, 
sclerotized connections both to juxta and base of costa of valva; juxta 
appearing to have a large, pointed, conical or thornlike medial process 
adjoining its posterior margin, seemingly apposed to end of aedeagus, 
and derivation of this structure also unclear. Vesica with clumps of 
small cornuti. 

FEMALE (Fig. 14, type species only): Head: Antenna simple, slen- 
der, sparsely setose, with scape much smaller than that of male. Labial 
palpus small, not exceeding front. Eye nearly as large as that of male. 
Ocellus absent. Tongue absent. Front, chaetosema, and legs similar to 
those of male. Brachypterous; forewing 6-7 mm long, % to % length 
of body, narrow, straplike, tapering to a pointed end, roughly clothed 


VOLUME 48, NUMBER 1 1 


with scales narrower than those of male and with both margins fringed 
with long, straight, bristlelike setae slightly longer than width of wing; 
hindwing present as small vestige beneath base of forewing and bearing 
long setae distally. Venation (Fig. 12) very reduced but bearing three 
longitudinal elements that are probably the stems of Sc, R, and Cu. 

Female genitalia (Fig. 13). Simple and without very significant fea- 
tures except an extremely long ostial cavity, with a pair of lateral 
sclerotized supports at base or in what may be a funnel-like posterior 
section of the ductus bursae, that on right side the larger; corpus bursae 
ovoid, membranous, delicate, without signum; ductus seminalis arising 
from bursa near ductus bursae. Sclerotized parts darkly pigmented. 

Distribution. Known only from arid habitats of eastern California, 
and western Nevada. 

Early stages. Eggs and first instar larvae were obtained, but larvae 
would not feed. Food plant unknown. Young larva typically geometroid, 
with no indication of a third pair of prolegs such as might indicate a 
relationship to the Alsophila group. 

Remarks. Sattler (1991) has reviewed wing reduction in Lepidoptera 
and analyzed implications of flightlessness. Examples of brachyptery 
in female winter moths in the Northern Hemisphere are known in 
several families; they are particularly numerous in Geometridae (e.g., 
Alsophila, Oenochrominae; Phigalia, Ennominae; and Operophtera, 
Larentiinae). The taxonomic relationships of Tescalsia proved difficult 
to determine and are still not clear. A combination of such features as 
the double accessory cell, free subcosta, extremely long fusion of Sc + 
Rs in the hindwing, and condition of the anal veins, points to a probable 
connection with the Larentiinae. Because of the elongate discal cells, 
loss of the proboscis, and reduced tympanic cavities, the possibility of 
relationship to the Alsophila group (currently in the Oenochrominae 
but probably misplaced) was also considered. All species of Alsophila, 
as well as the similar Phthorarcha primigera Staudinger (Central Asia) 
and Inurois tenuis Butler (Japan), were examined. These agree with 
one another in venation, especially with respect to the anal veins of the 
hindwing consisting of a weak Ist anal fold and well-developed 2nd 
and 3rd anals. Tescalsia clearly differs in having a strong Ist anal fold, 
well-developed 2nd anal, but no 3rd anal. This would seem to relate 
Tescalsia to the Larentiinae, in which the Hydriomenini and Operoph- 
terini have anal veins of this type. Surprisingly, Phthorarcha has wide, 
rounded wing scales almost exactly like those of Tescalsia, although 
other members of the Alsophila group and Larentiinae examined do 
not. 

The venation agrees best with that of the Larentiinae, although it is 
of an exaggerated type with two large accessory cells, elongated discal 


12 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


cells, and elongated fusion of Sc and Rs in the hindwing, and the almost 
straight, transverse closure of the ends of the cells is unusual. The 
genitalia, both male and female, seem closer to those of Operophtera 
than anything else, the male genitalia are quite like those of Operophte- 
ra in general form, in the shape of the valvae, and in the apparent 
homology of all components of the anellus. In female Operophtera, the 
beginnings of the same kind of large ostial cavity and short, sclerotized 
ductus bursae are apparent, and the simple, membranous corpus bursae, 
lacking a signum, agrees exactly. Operophtera has fairly well-developed 
tympanic cavities in the base of the abdomen, but their loss, together 
with the loss of the tongue, extremely wide pectinate male antennae, 
and curiously specialized female wings are adaptations to an extreme 
habitat. The large foretibial claw appears here and there in many groups 
of geometrids and noctuids, especially of desert habitats, and it has no 
important systematic significance. A palearctic geometrid that occurs 
in winter in Turkmenia and Kazakhstan, Cheimoptera pennigera Danil. 
(Danilevskiy 1969) shows many of the same features, including the 
foretibial claw and loss of the tympanic cavities, although it is unrelated 
to Tescalsia and believed to belong to the Ennominae. Tescalsia should, 
for the present, be assigned to the tribe Operophterini of the Laren- 
tiinae, although there are notable differences. For example, other species 
of Operophterini possess a reduced proboscis, large chaetosemata that 
meet in the middle in some instances, coarsely ciliate rather than bi- 
pectinate antennae in the male, lack the foretibial claw, and have either 
one accessory cell in the forewing (Operopthtera), or two (Epirrita) as 
in Tescalsia. 

The superficially similar, gray, long-winged Chesiadodes morosata 
Hulst (Ennominae) flies in the same area near Lone Pine, Inyo County, 
California, in December, and also has a foretibial claw, although of 
different shape. It differs in having a proboscis. The two are not closely 
related, and the female of Chesiadodes has fully developed wings. 


Tescalsia giulianiata Ferguson, new species 


(Figs. 1, 2, 5-9, 11-14) 


Diagnosis. Males large, forewing length 16.0-17.5 mm; wings gray 
with blackish markings; hindtibia with two pairs of spurs. Female as 
described for genus and illustrated. 

Description. MALE: Head: labial palpus small, not surpassing front; 
eye of about same dimensions as that of T. minata but more protuberant, 
its form exceeding that of half a sphere; front bulging, roundly convex, 
with large, broad, gray-brown or whitish scales, tending to be oriented 
toward middle of front; eye rimmed anteriorly and ventrally with 


VOLUME 48, NUMBER 1 13 


Fics. 1-7. 1, Tescalsia Pliaihata Eur dteoe holstene male. 2, T. Balaniata para- 
type female sare data as holotype). 3, T. minata Ferguson, felon ee male; 4, T. minata, 
paratype male. 5-7, T. giulianiata, female (Deep Spring Valley, CA, XII-15-78): 5, 
perched on sand; 6, in repose balanced on wingtips; 7, climbing on Chenopodium branch. 


contrastingly pale border of radiating whitish scales. Thorax beneath 
without long, hairlike scales; legs similar to those of T. minata except 
that femora lack long, hairlike scales, and hindtibia has two pairs of 
spurs. Wings whitish, dusted with blackish scales and thus appearing 
gray, although forewing sometimes so heavily suffused that markings 
are obscured; forewing normally with diffuse, slightly dentate or sinuous 


14 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


dark antemedial and postmedial bands, the former slightly convex and 
subparallel to postmedial, which is nearly parallel to outer margin; 
subterminal shade, if present, indistinct, similarly parallel to outer mar- 
gin; some veins, especially in medial area, faintly outlined with dark 
scales; black discal spot present; fringe white, checkered with dark 
brown and preceded by weak terminal line of diffuse dark spots. Hind- 
wing paler gray, almost unmarked except for small discal spot and 
diffuse but complete transverse band crossing just beyond middle of 
wing; fringe whitish, unmarked, preceded by faint, broken terminal 
line in some specimens. Underside with fore- and hindwing nearly 
alike, light gray with discal spots, diffuse postmedial bands, and variable, 
black, interrupted, terminal lines. Length of forewing: holotype, 17.0 
mm; other 6, 16.0-17.5 mm. Genitalia (Fig. 11), differing from those 
of T. minata mainly in their wider, blunt or obtusely pointed medial 
juxtal process, less heavily spined manica, and smaller, paired, setose 
processes attached to bases of costal sclerite. Eighth sternite rectangular, 
not triangular, and eighth tergum without a posterior border of per- 
sistent scales. 

FEMALE: As described for genus. Antenna simple, covered with 
whitish scales. Hindtibia with two pairs of spurs. Body gray, variably 
dusted with black scales; in paler specimens a pair of dark subdorsal 
spots toward posterior margins of abdominal segments 2-5. Legs gray 
brown with each tarsal segment pale-tipped. Forewing with light and 
dark scales almost evenly mixed and no other markings. Length of 
forewing: 5-6 mm. Genitalia (Fig. 18) as illustrated and described for 
genus. 

Types. Holotype male and allotype female: CALIFORNIA, Inyo Co., 
Alabama Hills, 4 miles [5 airline km] southwest of Lone Pine, 4550’ 
[1390 m elev.], 6/7-XII-1982, in pitfall traps (J. De Benedictis & J. 
Powell); deposited in Essig Museum of Entomology, U. California, 
Berkeley. Paratypes (15), all Inyo Co., CA: 44, 3 2, same data as holotype, 
7 /11-XII-1982 (D. Giuliani, De Benedictis, Powell); 1 6, Alabama Hills, 
N. fork Lubkin Cr., 4 mi. S, 1 mi. W of Lone Pine, 4800’, 11-XII-1982 
(Giuliani); 1 2, Deep Spring Valley, 8-XII-1978, sand dunes (Giuliani); 
2 2, same locality, 15/16-XII-1978, pitfall traps (Powell); 1 6, same 
locality, 17-XI-1980, flying at sundown (Giuliani); 2 6, same data, 17- 
XI-1982; 1 2, Owens Lake, mid XI to mid XII-1978, ethylene glycol 
pit trap, Atriplex-Franseria assoc. (Giuliani, F. Andrews, D. Hardy); 
paratypes deposited in California Dept. Food & Agric., Sacramento, 
Essig Museum, and U.S. National Museum of Natural History, Wash- 
ington, D.C. 

Habitats. Tescalsia giulianiata is known from three sites in the Owens 
Valley region, California, which are similar in general aspects of veg- 


VOLUME 48, NUMBER 1 15 


sl mee 


8 


Fic. 8. Tescalsia giulianiata Ferguson, male, wing venation. 


etation architecture, with a low scrub of scattered shrubs interspersed 
with open patches of sand, yet they differ in dominant plant species. 

1) Deep Spring Valley is a closed basin at 1500-1600 m elevation, 
situated southeast of the White Mountains and surrounded by arid 


16 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


mountain ridges. To the west the valley is separated from the Owens 
Valley by Westgard Pass (2225 m) and to the east from the valleys of 
Nevada by Gilbert Summit (1950 m). From the highway along the 
north edge of the valley, the terrain slopes southward towards Deep 
Spring Lake, which is dry except following winter storms. Fine aeolian 
sand from the dry lakebed has been deposited to form low ridges and 
dunes that are stabilized by low scrub dominated by Atriplex confer- 
tifolia (Chenopodiaceae), Thamnosma montana (Rutaceae) and scat- 
tered patches of Chrysothamnus nauseosus (Asteraceae). Female moths 
were taken from open sand and in pitfall traps placed at the base of 
Atriplex and Thamnosma, about 1 km north of the lakebed. 

The valley is about 300 m above the floor of Owens Valley, and it 
evidently acts as a basin for cold air drainage, with snow patches per- 
sisting much longer than at comparable elevations on the hills around 
Owens Valley. During our December visits, daytime temperatures of 
10-18°C fell rapidly, to 4.5-5.0°C towards sundown and 1.6-4.5° by 
dusk. 

2) The Alabama Hills are remnants of an ancient uplift that has 
eroded to low hills of decomposed granitic, alluvial sand subtending 
weathered, granite outcrops. The collecting site is situated at 1890 m 
elevation about 5 airline km southwest of Lone Pine, off Indian Springs 
Road. This area is characterized by gently sloping expanses of coarse, 
granitic sand stabilized by a low scrub consisting mainly of Tetradymia 
glabrata (Asteraceae), a low spiny Atriplex, and scattered Chrysotham- 
nus nauseosus. Most of the type series was taken at this site, December 
7/11, 1982, in pitfall traps deployed December 6. Although the locality 
is only about 100 m lower than Deep Spring Valley, it isa much warmer 
habitat during winter. 

3) The former Owens Lake was drained by the Los Angeles water 
district, beginning in the 1920’s, and has long been a dry lakebed. 
Aeolian sand is deposited in low ridges along its east margin. One female 
of T. giulianiata was taken here in a pitfall trap. Giuliani and F. G. 
Andrews had monitored the area by pitfall trapping, with 12 traps in 
each of 7 vegetation types for one year. The site that yielded the 
Tescalsia is about 3 km northwest of Keeler (1100 m). The traps were 
set east of a few low sandhills at the margin of the lakebed, in a low 
area that acts as a catch basin for the sparse rainfall runoff. It is char- 
acterized by shadscale scrub, alkaline tolerant species of Chenopodia- 
ceae, including Atriplex hymenelytra, the spiny Atriplex of the Ala- 
bama Hills, Allenrolfia occidentalis, and Sarcobatus vermiculatus, as 
well as Tetradymia glabrata. Thus the dominant vegetation is more 
similar to the Alabama Hills site than to Deep Spring Valley. We did 
not deploy additional pitfall traps at this locality after. 1978. 


VOLUME 48, NUMBER 1 17 


Diel rhythm. Temperatures in December in the Owens Valley area 
typically drop to 2-10°C below freezing at night but rise well above 
freezing during the day, often to 10-15°C. Nonetheless, our observations 
suggest that Tescalsia giulianiata is not diurnal and begins activity near 
sundown. Its period of activity may vary daily with a low temperature 
threshold and may exclude some evenings. Males were observed flying 
only twice, by Giuliani on November 17, 1980 and Nov. 17, 1982, near 
sundown. One female was found on open sand in late afternoon, and 
one was trapped in a pitfall between 1530-1615 PST (ca. 8—10°C; sunset 
at 1610), and another between 1630 and 1030 PST the following day, 
in temperatures below 5°C. Other individuals were trapped during 
longer intervals, including one male and a female at the Alabama Hills 
between 1230 and 0930 the following day. Mating was not observed. 

On each of the four dates in December, 1977-82, we made continuous 
observations for 2-8 h, on days when temperatures rose from 0.5—2.7°C 
at 0930-1030 PST to 15°C in mid afternoon, dropping to 4.5-5.0°C by 
sundown. No males were seen, and just one female was trapped during 
these intervals, which included extensive pitfall trapping, beating of 
shrubs, sifting and net brushing of the sand. T. giulianiata evidently 
is not adapted to diurnal flight when temperatures are highest. We ran 
a blacklight trap just once; the temperature was 4.5°C at dusk, and not 
one insect was trapped. 

One captive female survived more than 4 days, with lab temperatures 
at 10-14°C at night to 21°C diurnally. Activity periods were sporadic, 
but in absence of disturbance, she seemed most active during evening, 
1800-2100 h, moved only slowly when viewed by flashlight at 0500- 
0700, and rarely during morning hours. 

During periods of activity, the female held the antennae and fore- 
wings at about a 45° angle to the plane of the body (Figs. 5-7). When 
quiescent, she positioned the antennae back along the body, and the 
wings were curled downward. On the sand surface, she seemed very 
awkward, scarcely able to ambulate forward or walk evenly. Walking 
on the sand, she dragged her abdomen, leaving linear tracks, but this 
did not seem to be a part of oviposition behavior. By contrast, when 
debris or the branchlet was encountered, she climbed quickly with agile 
movement from twig to twig and often hanging by one or two legs, 
reminiscent of a miniature orangutan. She frequently perched on the 
highest reach of the branchlet and moved quickly to maintain that 
position if the branch was moved. It seems that the species is adapted 
to life in shrubs rather than on the sand, which fails to explain why 
females were trapped in pitfalls but none could be beaten from shrubs. 
In fact, there may be strong selection against life on the sand at that 
time of year because predaceous mites and lycosid spiders appeared in 


18 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fics. 9-11. 9, Tescalsia giulianiata Ferguson, female, right prothoracic leg; small 
process arising near middle of double clawed tibia is a vestigial epiphysis. 10, T. minata 
Ferguson, male genitalia; a, aedeagus. 11, T. giulianiata, male genitalia; a, aedeagus. 


VOLUME 48, NUMBER 1 19 


33-50% of our pitfall traps. Two of the female T. giulianiata fell victim 
to these predators overnight, when we used dry cups. 

Whether on sand or on twigs, the fore tarsi were extended, so that 
the tibial hooks were well back from the substrate; evidently they are 
not employed in adult locomotion. Upon disturbance, the female con- 
sistently feigned death, falling on her side with all legs retracted, and 
remained so for 10—20 seconds. Also, at times the female assumed a 
repose stance, balancing on her wingtips and curled abdomen, with the 
legs partially folded (Fig. 6). This may have been an abnormal behavior 
in confinement, but it was repeated several times for lengthy periods. 

Oviposition. The single female retained alive was housed in a plastic 
box with sand and a dry branch of Atriplex (JAP 78M1). A cotton wick 
was provided, but there was no indication that the female ever imbibed 
moisture or touched the cotton with her vestigial mouthparts. She de- 
posited 10 eggs in captivity, none during the first 48 h, 7 in the 8rd 
24-h period, and 3 the following day. Eggs were deposited between 
0940-1000 PST (1), 1055-1125 (1), 1400-1700 (6), and one later. Most 
were placed on loose sand, but the female was not observed to probe 
the sand with her abdomen. Two eggs were deposited on the damp 
cotton wick after the female had been coaxed onto it, and one egg was 
affixed to a dry Atriplex leaf. 

Cuming (1961) reported the activity periods of the related larentiine 
winter moth, Operophtera brumata (L.), to be nocturnal. Caged moths 
in outdoor conditions mated between 1730-2305 at temperatures of 
—0.8 to +12.2°C, and females oviposited between 1515-0020 at —1.6 
to +12.2°C. T. giulianiata displays a comparable diel rhythm and 
temperature tolerance. 

Eggs. The eggs (n = 10), which were deposited singly, were oblong, 
slightly variable in shape, 0.80-0.90 mm long x 0.60 mm wide x 0.48 
mm thick, somewhat flattened on the side away from the substrate. 
The chorion was opaque, white, very weakly rugose, appearing smooth 
under low magnification. When first deposited the eggs were pale 
greenish, similar in color and shape to new leaves of Atriplex confer- 
tifolia. 

The eggs were confined in individual gelatin capsules; five were 
placed in a tightly covered plastic container in refrigeration each night 
(ca. 8:16 h, 1.6°: 15-20°C), while the remainder were retained at lab 
temperatures (10—20°C), for 30 days. During development, eggs dark- 
ened and most showed a depressed area away from the micropylar end 
as though collapsing. Larvae began eclosing in the refrigerated sublot 
January 21, after 34 days; the others were refrigerated 8:16 h from day 
36 to 46, and first instar larvae emerged February 2-4, after 46-48 
days. 


20 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


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S 


Md 


Fics. 12-18. 12, Tescalsia giulianiata Ferguson, female, forewing showing setation 
of margins and rudimentary venation. 13, T. giulianiata, female genitalia. 


First instar larvae. The eclosing larva ate a ragged, round hole at 
the micropylar end but did not continue to feed on the chorion. Each 
was placed in a small cup with synthetic diet (modified Shorey Diet 
used for Choristoneura) and a sprig of fresh Atriplex (but not A. con- 
fertifolia) from the U. C. Botanic Garden. None fed. After two days, 


VOLUME 48, NUMBER 1 | WAI 


\ Wy 
Nur 


ja 


\\) 


YR 


aS aS 
= aw 


SQ Sant A 
i 


=e ww SS 


Fic. 14. Adult female of Tescalsia giulianiata. 


some larvae also were offered a sprig of Prunus or Salix, but by that 
time they likely were too weak to feed. All died by the 3rd day after 
eclosion. 

If failure to accept synthetic diet and the plants provided indicates 
a narrow host specificity, we cannot suggest a probably food plant. In 
retrospect, after observing the three habitats from which the flightless 
females were taken (just the one site was known at the time the eggs 
were obtained), Atriplex is the most plausible of the dominant, woody 
shrubs, with one or more species occurring in all three habitats. Chrys- 
othamnus, which was present only in scattered patches, or some her- 
baceous perennial are possible alternative candidates. 


Tescalsia minata, Ferguson, new species 


(Figs. 3, 4, 10) 


Diagnosis. Males small, forewing length 12-15 mm; wings brown 
with mostly indistinct, darker brown markings; hindtibia with one pair 
of spurs. Female unknown. 


22 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Description. MALE: Head: labial palpus slender and delicate, but 
with its terminal scales clearly surpassing front; eye about as in the 
other species but much less protuberant, its shape distinctly less than 
that of half a sphere; front somewhat protuberant but flattened rather 
than roundly convex, with large, broad scales tending to be oriented 
toward middle of front; eye rimmed anteriorly and ventrally with 
brown scales concolorous with those of front, not contrasting. Thorax 
beneath and tegulae above sparsely clothed with long, brown, hairlike 
scales; legs similar in the two species except that femora of T. minata 
have long, hairlike scales, and the hindtibia only one pair of spurs. 
Wings gray brown with variable, darker brown markings; forewing 
with antemedial band often strongly convex and enclosing a paler area 
toward base; faint basal band also may be present; postmedial not always 
distinct but, if present, not parallel to outer margin, more curved, 
concave in posterior half of wing and convex in costal half; a vague, 
pale, subterminal band may be present; dark-brown discal spot present 
but weak; fringe brown, concolorous with wing, unmarked; terminal 
line wanting. Hindwing brown, hardly paler than forewing, unmarked 
except for weak discal spot; fringes concolorous. Underside much like 
upperside except that a faint, diffuse, convex, transverse band may cross 
just beyond discal spot, and veins on underside of hindwing may in 
part be faintly outlined with darker brown scales. Length of forewing: 
holotype, 18 mm; other 6, 12-15 mm. Genitalia (Fig. 10), similar to 
those of T. giulianiata; the most obvious difference is in the shape of 
the large medial process of juxta. In T. minata it has an abruptly 
acuminate, thornlike shape, with a sharp-pointed apex. Spines on man- 
ica more numerous and larger, and paired, setose processes flanking 
manica and adjoining base of costal sclerite of valva are larger. Eighth 
sternite small and triangular rather than quadrate, and eighth tergum 
with wide, dense tuft of short, persistent scales along its posterior mar- 
gin, not easily removed in dissection. 

FEMALE: Unknown. 
Types. Holotype 6, Mina [Mineral County], Nevada, November 17, 
1914, A. Wetmore. Paratypes: 4 4, same data. Type series in collection 

of U.S. National Museum of Natural History, Washington, D.C. 


ACKNOWLEDGMENTS 


The discovery of this genus and subsequent important collections were made by Derham 
Giuliani, of Big Pine, California, who has contributed considerably to our knowledge of 
the insects of the interior deserts in California. F. G. Andrews, California State Department 
of Food & Agriculture, Sacramento, provided support for pitfall trap sampling by Giuliani. 
J. A. De Benedictis, now at University of California, Davis, J. T. Doyen, J. K. Liebherr, 
now at Cornell University, Ithaca, New York, endured winter field trips while at Berkeley. 
The genitalia and wing venation drawings were done by Linda H. Lawrence, Staff 


VOLUME 48, NUMBER 1 23 


Illustrator, Systematic Entomology Laboratory; the habitus drawing by Tina Jordan, U. 
California, Berkeley. The photographs of specimens were made by Ferguson and those 
of living moths by Powell. 


LITERATURE CITED 


CUMING, F. G. 1961. The distribution, life history, and economic importance of the 
winter moth, Operophtera brumata (L.) (Lepidoptera, Geometridae) in Nova Scotia. 
Canad. Entomol. 93:1385-142. 

DANILEVSKI, A. S. 1969. Two remarkable new species of winter moths from the deserts 
of Soviet Central Asia: Dasyethmia hiemalis, gen. et sp. n. (Lepidoptera, Ethmiidae), 
and Cheimoptera pennigera, gen. et sp. n. (Lepidoptera, Geometridae). Entomol. 
Rev., Washington, D.C. 48:176-191. 

SATTLER, K. 1991. A review of wing reduction in Lepidoptera. Bull. Brit. Mus. Nat. 
Hist. (Entomol.) 60:243-288. 


Received for publication 5 June 1993; accepted 31 July 1993. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 24-45 


SPLIT SKIPPERS: MEXICAN GENUS POANOPSIS GOES 
IN THE ORIGENES GROUP—AND YVRETTA FORMS 
THE RHESUS GROUP—OF POLITES (HESPERIIDAE) 


JOHN M. BuRNS 


Department of Entomology, National Museum of Natural History, 
Smithsonian Institution, Washington, District of Columbia 20560, USA 


ABSTRACT. Genitalia show that the montane Mexican genus Poanopsis Godman 
fits snugly within the origenes group of Polites Scudder and that Yoretta Hemming 
constitutes a new species group of Polites. Relationships have been masked in part by 
evolutionary reduction and loss of various characters. The origenes group, with the 
northern superspecies Polites mystic (P. mystic (Edwards) + P. sonora (Scudder)), the 
intermediate linking species P. origenes (Fabricius), and the southern sister species P. 
puxillius (Mabille) new combination and P. pupillus (Plétz) new combination, extends 
from the width of southern Canada to southern Mexico. The rhesus group of Polites, 
with P. rhesus (Edwards) and the sister species P. carus (Edwards) and P. subreticulata 
(Pl6tz) (new combinations, all), extends from southern central Canada to Panama. Geo- 
graphic distributions of species within these groups (and in the themistocles and vibex 
groups of Polites, as well) still strongly reflect allopatric speciation. 


Additional key words: genitalia (male and female), variation, generic limits, character 
reduction and loss, speciation (allopatric). 


In dealing with biodiversity, systematists divide and conquer. For- 
mally described divisions assume a life of their own. The longer they 
live and the more we use them, the sounder they seem. Sometimes we 
get so accustomed to overly fine divisions that we cannot see the forest 
for the trees. The American genus Polites Scudder is one such forest, 
a good bit larger and denser than we thought. 

Once upon a time, Godman (1900) created Poanopsis for a small, 
brown, high montane, Mexican skipper, Pamphila puxillius Mabille, 
which he made the type of his monotypic new genus. He observed that 
puxillius, with its short, broad wings, looks like massasoit Scudder, the 
type of the genus Poanes Scudder (hence the name Poanopsis), but 
differs conspicuously in having an oblique stigma on the forewing of 
the male from the origin of vein 3 to vein 1. Godman (1900:pl. 98, fig. 
40—my Fig. 1) illustrated the male genitalia of Poanopsis puxillius 
without comment. In mid century, Evans (1955) added a similar Mex- 
ican skipper, pupillus Plotz (=catahorma Dyar), to Poanopsis, carica- 
tured the male genitalia of both species, and set Poanopsis next to 
Poanes, eight genera removed from Polites. However, genitalia show 
that both species of Poanopsis belong in one of the species groups of 
Polites. 

Polites is a familiar, widespread, New World genus, stretching from 
Canada to central Argentina, from Atlantic to Pacific coasts in both 
Americas, and through much of the West Indies. On the basis of genitalic 


VOLUME 48, NUMBER 1 25 


Fic. 1. Male genitalia of Polites puxillius (minus left valva) in left lateral view (ex 
Godman 1900:pl. 93, fig. 40). This figure presents the inner surface of the right valva 
whereas those that follow present the outer surface of the left valva. The specimen is 
“from the high tablelands of MEXICO.” 


and stigmal characters, MacNeill (1993) recognized four subdivisions: 
the themistocles group, the origenes group, the vibex group, and Polites 
baracoa (Lucas). He pointed out that, within groups (the themistocles 
group in particular), genitalia may be so conservative and so individ- 
ually variable that differences between species blur but that, paradox- 
ically, superficial color pattern will distinguish some of the genitalic 
look-alikes. 


THE ORIGENES GROUP OF POLITES 


The origenes group contains the eastern and central North American 
Polites origenes (Fabricius) plus the continent-spanning and largely 
allopatric sisters P. mystic (Edwards) and P. sonora (Scudder), all of 
which show some internal differentiation. From an evolutionary per- 
spective, P. mystic and P. sonora can be considered a superspecies (see 
Stanford & Opler 1993:74, 75 for maps that more or less reflect the 
mutual geographic replacement of these two similar species, and see 
Burns 1964, 1983 for detailed analyses of superspecies in other American 
skippers). The origenes group ranges from Canada to the southern 
United States and, disjunctly, to higher elevations in the Sierra San 
Pedro Martir of Baja California Norte, Mexico. Now, with Polites pux- 
illius new combination and Polites pupillus new combination, it jumps 
from three to five obvious species and from the southern United States 
to southern mainland Mexico. Though these five species vary greatly 
in facies, they are genitalically close, especially in males. 

To be sure, male genitalia are grossly similar throughout the genus 
Polites (see figures in Scudder 1889, Skinner & Williams 1924 [or Lind- 
sey et al. 1931], MacNeill 1993). Of all parts, the aedeagus is most 
generally useful in clustering species. Its accessories at once stamp the 
origenes group: the paired, toothed titillators are large, boldly dentate 
plates, something like semicircular saws (Figs. 1-5), extending down- 
ward from the lower part of the flared, distal end of the aedeagus (Figs. 
1, 3, 5); and the lone cornutus is a rolled scroll bearing a crown of thorns 
(Figs. 2-5). In all other Polites the paired, toothed titillators assume 


26 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 2,3. Male genitalia of Polites puxillius from 28 mi (45 km) E EI Salto, 8400 ft 
(2560 m), Durango, MEXICO, 6-7 August 1972, Viers & MacNeill [genitalia dissection 
number X-2652] (USNM). Scale = 1.0 mm. 2, Tegumen, uncus, tip of gnathos, both 
valvae, both titillators (each with four teeth), and the single cornutus (like a rolled scroll 
bearing a crown of thorns)—all in posterior view; 3, Complete genitalia (minus right 
valva, right titillator, and juxta) in left lateral view, with vesica everted. 


— 


Fics. 4,5. Male genitalia of Polites pupillus from Sierra de Guerrero, MEXICO, July 
1913, R. Miller [X-2699] (USNM) (type of Amblyscirtes catahorma Dyar). Scale = 1.0 
mm. 4, Tegumen, uncus, tip of gnathos, both valvae, both titillators (left with 10 teeth, 
right with 8), and the single cornutus (like a rolled scroll bearing a crown of thorns)— _ 
all in posterior view; 5, Complete genitalia (minus right valva, right titillator, and juxta) 
in left lateral view, with vesica everted. 


VOLUME 48, NUMBER 1 Daye 


28 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


very different shapes, come from the upper (rather than lower) part of 
the distal end of the aedeagus, and, if they extend anywhere, extend 
backward (not downward); decorated cornuti number two or three 
(instead of one) and suggest thorny scrolls only in the vibex group (where 
the aedeagus sports a unique, midventral, caudally-directed prong). 
The distal end of the uncus, which is divided, is bent farther dorsad 
in the origenes group (Figs. 1, 3, 5) than in any other group of Polites. 
Female genitalia in Polites present less of a generic gestalt, varying 
more both within and between groups—and, to some degree, in par- 
allel—so that group characterization is not as simple. In the origenes 
group (also in the vibex group, but in no others) the apophyses anteriores 
join the lamella postvaginalis via continuously sclerotized bands (Figs. 
6-9). In the origenes group (also in P. baracoa) sclerotization of the 
lamella postvaginalis is intermediate (P. puxillius [Figs. 6, 7], P. pupillus 
[Figs. 8, 9], and P. origenes) to extensive (superspecies P. mystic) rather 
than limited (as it is in the themistocles group [MacNeill 1993] and in 
the vibex group [where the lamella postvaginalis carries a unique, mid- 
ventral, short, sclerotized, finely spined keel]). In the origenes group 
sclerotization of the ductus bursae is partial and posterior; but it may 
be ventral only (superspecies P. mystic), ventral and lateral (P. pupillus 
[Figs. 8, 9]), or ventral, lateral, and, in a very narrow band, dorsal (P. 
puxillius [Figs. 6, 7], P. origenes, and, at times, vaguely, P. pupillus)— 
always with a longitudinal, midventral groove (Figs. 6-9) or break in 
the sclerotization. (Sclerotization of the ductus bursae is partial, pos- 
terior, but 360° and broadly ringlike [though midventrally weak] in the 
vibex group; partial, lengthy, mostly ventral but also somewhat lateral 
[resembling a long, essentially ungrooved, scoop] in P. baracoa; and 
complete, extending more or less the entire length of the ductus bursae 
and running all the way around, usually with a strong middorsal groove 
and a ventral pouch, in the themistocles group [MacNeill 1998].) 
While the northern major differentiates of the origenes group of 
Polites (P. origenes and superspecies P. mystic) are widely known, 
repeatedly shown in the rising tide of North American butterfly books, 
and readily separable superficially, the southern differentiates (P. pux- 
illius and P. pupillus) are not and must be treated here. Whereas in 
both sexes of mystic and sonora and in males of origenes extensive 
yellow to yellow-orange brightens the dorsal wing surface, in females 
of origenes and in both sexes of puvxillius (Figs. 23, 24) and pupillus 
(Figs. 25, 26) the wings above look basically brown (with a variable set 
of small, pale spots [yellowish in origenes, white in puxillius and pupil- 
lus]). Wingshape is sexually dimorphic in the northern three species 
but not in the southern two: in mystic, sonora, and origenes the wings 
of males are narrower and more pointed, those of females, broader and 


VOLUME 48, NUMBER | 29 


rounder; but in puxillius and pupillus the wings are broad and round 
in both sexes (Figs. 23-26). So, with respect to both color and wingshape, 
puxillius and pupillus tend to resemble females of origenes. 

In the two Mexican species, wing spots, which are white, are expressed 
better by pupillus (Figs. 25, 26) than by puvxillius (Figs. 23, 24). Fore- 
wing spots of pupillus look whiter and brighter partly because, in some 
to most of them, some scales stand up from the surface of the wing so 
as to let light through. In puvxillius all the white scales lie flat against 
the wing so that all spots are opaque, none hyaline. Usually, pupillus 
develops a full set of forewing spots—in spaces 1b, 2, 3, 4, 5, 6, 7, 8, 
and the cell (Figs. 25, 26)—but puxillius rarely does, almost always 
dropping the one or two spots in space 1b (Figs. 23, 24), often skipping 
those in spaces 4 and 5, as well as that in 8 (Fig. 23), and, in one male 
examined, losing all but spots 3 and 6. At least some hindwing spots 
(which are opaque) appear dorsally in pupillus (usually spots 2 to 6) 
(Figs. 25, 26) while none really do in puxillius (Fig. 23) (at most, there 
may be a suggestion of a spot in space 3—see Fig. 24). Ventrally, where 
spot development is better, up to a full set may surface in pupillus— 
in spaces lc, 2, 3, 4, 5, 6, 7 (rare) and the cell (Figs. 25, 26); in puxillius 
ventral expression runs the gamut from all except the rare spot 7 (Fig. 
24) to nothing whatsoever. 

As for real interspecific differences in the conservative but individ- 
ually variable male genitalia, titillator teeth are fewer in puxillius (Figs. 
1-3), varying from 4 to 6 (usually 4) on each of the paired titillators, 
and commoner in pupillus (Figs. 4, 5), where they vary from 6 to 10 
(usually 7 or 8). The valva of puxillius, in lateral view (Fig. 3), is more 
tapered distally along its ventral margin and hence not as broad (from 
top to bottom) at its distal end as it is in pupillus (Fig. 5). (Evans does 
not mention these differences; yet once you know what to look for, you 
can detect both of them in his [1955:pl. 78, M.21.1, M.21.2] genitalic 
cartoons.) In posterior view, the lower part of the divided distal end of 
the valva usually forms more of a U in puxillius (Fig. 2) than it does 
in pupillus (Fig. 4). Though conspicuous in the specimens figured, this 
difference almost vanishes when certain variants are compared. The 
greater valval height of pupillus, evident in lateral view, usually shows 
too in posterior view as a ventral, vertical extension below the U (com- 
pare Fig. 4 with Fig. 2). 

One interspecific difference in the highly variable female genitalia 
crept into the foregoing group comparisons: the incomplete scleroti- 
zation of the ductus bursae (mainly ventral and lateral) continues dor- 
sally to form a narrow but unmistakable sclerotized ring around the 
ductus in puxillius (Figs. 6, 7) but not—or, at best, very weakly—in 
pupillus (Figs. 8, 9). In addition, the sides of the midventral groove in 


30 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 6, 7. Female genitalia of Polites puxillius from 25 mi (40 km) W Durango, 
Durango, MEXICO, 5 August 1972, MacNeill & Viers [X-2653] (USNM). Scale = 1.0 
mm. 6, Ovipositor lobes (retracted), eighth tergite with apophyses anteriores (sclerotically 
connected to the lamella postvaginalis of the sterigma), sterigma, and bursa copulatrix in 
ventral view; 7, The same, plus the right apophysis posterioris and part of the ductus 
seminalis, in right lateral view. 


the ductal sclerotization are more divergent anteriorly in puvxillius (Fig. 
6) than in pupillus (Fig. 8), and the sclerotization itself is more fluted 
in puxillius (Figs. 6, 7) than it is in pupillus (Figs. 8, 9). The lateral 
part of the ductal sclerotization (which is more extensive on the left 
side than on the right—see Figs. 7, 9) is more extensive on both sides 
in pupillus than it is in puxillius; so it better hides the roof of the 


VOLUME 48, NUMBER 1 31 


Fics. 8,9. Female genitalia of Polites pupillus from 34 mi (55 km) SE Acatlan, 6000 
ft (1830 m), Puebla, MEXICO, 9 July 1952, E. E. Gilbert, C. D. MacNeill [X-3198] 
(MacNeill collection). Scale = 1.0 mm. 8, Ovipositor lobes (exserted) with apophyses 
posteriores, eighth tergite with apophyses anteriores (sclerotically connected to the lamella 
postvaginalis of the sterigma), sterigma, and bursa copulatrix in ventral view; 9, The 
same, plus part of the ductus seminalis, in right lateral view. 


32 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


midventral groove in lateral view in pupillus (Fig. 9) than in puxillius 
(Fig. 7). Again in lateral view, the roof of the midventral groove is 
convex in puxillius (Fig. 7) but straight in pupillus (Fig. 9). All these 
interspecific differences are rather subtle. 

Polites pupillus is larger than P. puxillius. In each species the fore- 
wing of the female averages about one mm longer than that of the 
male, and in each sex the forewing of pupillus averages about one mm 
longer than that of puxillius: 


SEX mean range n 
puxillius 3 13.6 12.6-14.4 9 
2 14.9 14.0-15.6 13 
pupillus 3 14.8 13.5-16.2 16 
2 ony 15.0-16.4 OD 


The number of segments in the nudum of the antenna ranges from 11 
to 13 in puxillius (mean = 12.2, n = 21) and from 11 to 14 in pupillus 
(mean = 12.8, n = 20). In both species nudum segments are evenly 
split between the body of the club and the apiculus. 

Although both species are montane, P. puxillius is higher, altitudi- 
nally tighter (recorded from 8000 to 8500 ft [2440-2590 m]), and less 
widely distributed (Durango [Fig. 10]—to as far south as Guerrero 
according to Godman 1900, Hoffmann 1941). Polites pupillus extends 
from 4200 to 7500 ft (1280-2285 m) and from Sonora and Sinaloa to 
Colima, Puebla, Guerrero, and Oaxaca (Fig. 10). So far as known, flight 
times are similar: dates on specimens of puxillius examined run from 
18 July to 11 August; on pupillus, from 3 July to 8 August. 


THE RHESUS GROUP OF POLITES 


The rhesus group of Polites comprises at least three species (usually 
treated as two) that collectively range from the grasslands of southern 
Saskatchewan and Alberta, Canada, through the western Great Plains, 
southern Rocky Mountains, and southwestern United States, through 
Mexico and Central America, to the middle of Panama. Although God- 
man (1900:474) noted that “the structure of the genitalia of the males 
is very similar in the two species’ when he put them in his new genus 
Chaerephon, he did not see the great similarity between their genitalia 
(Godman 1900:pl. 93, figs. 4 and 7—my Figs. 15, 16) and those of what 
he called Thymelicus vibex (pl. 93, fig. 14) and Poanopsis puxillius (pl. 
93, fig. 40—my Fig. 1), which were on the same plate and which now 
are both in Polites. Likewise, Skinner and Williams (1923) figured the 


VOLUME 48, NUMBER 1 33) 


\ 
i) 
i 
| 
\ 


1 
a, —™. 
Vay Sn ~L, 


400 MILES 


300 500 KILOMETERS 


Fic. 10. Geographic distribution (based on plottable material examined) of Mexican 
sister species Polites puxillius (triangles) and P. pupillus (dots). 


male genitalia of the two US. species of Chaerephon and then (1924) 
those of nine U.S. species of Polites without seeing how remarkably 
close they are. Enamored of names, Hemming (1935) saw that Chae- 
rephon Godman is a junior homonym and replaced it with Yvretta. 
Commenting that its “genitalia [are] very like those of Polites,’ Evans 
(1955:315) still set Yuretta next to Hylephila, six genera removed from 
Polites (and 21 genera from Poanopsis). MacNeill (1993:177) observed 
that “male genitalic similarities suggest that the nearest relatives of 
Polites are the genera Yuretta Hemming, Hylephila Billberg, and Wal- 
lengrenia Berg.” I am going much further by reducing Yoretta to a 
new species group of Polites, with the species Polites rhesus (Edwards), 
P. carus (Edwards), and P. subreticulata (Pl6tz) (new combinations). 

Like all other elements of the male genitalia, the aedeagus in the 
rhesus group has that Polites look: anteriorly narrow, it at least doubles 
in size (in lateral view) with the entry of the ductus ejaculatorius and 
more or less flares at the elaborate distal end, which flaunts a pair of 


34 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 11-18. Male genitalia of Polites subreticulata from Coatepec, Veracruz, MEX- 
ICO, October 1910, R. Miller [X-3499] (USNM). Scale = 1.0 mm. 11, Tegumen and 
uncus (gnathos hidden) in dorsal view; 12, Complete genitalia (minus right valva and 
juxta) in left lateral view (left titillator more dorsal and more tightly dentate than right), 
with vesica everted showing all three cornuti (two dentate [upper one curved, lower one 
straight] and one without teeth); this individual somewhat malformed anteriorly: saccus 
curved sharply upward, aedeagus bent to right, and bottom of tegumen not curved ventrad 
near juncture with vinculum (compare Fig. 14); 13, Aedeagus in dorsal view (left titillator 
more tightly dentate than right), with vesica oieted (only the two dentate cornuti visible). 


toothed titillators as well as toothed cornuti in the vesica (Figs. kj 3s0% 
12-16, and figs. 11-19 in MacNeill 1993). 

In the rhesus group, the paired, toothed titillators come from the 
upper part of the distal end of the aedeagus, extend backward with 


VOLUME 48, NUMBER 1 395 


Fic. 14. Male genitalia (digital image) of Polites rhesus from Sapillo Creek Valley, 
5800 ft (1770 m), 12 mi (19 km) N Pinos Altos, Grant Co., New Mexico, USA, 28 May 
1959, J. M. & S. N. Burns [X-2712] (USNM). Complete genitalia in left lateral view (right 
titillator out of focus and toothless cornutus hidden), with vesica everted showing both 
dentate cornuti (upper one curved, lower one straight). 


one twist along their narrow length, and then expand distally into the 
dentate portion, with 3 to 7 teeth (usually 5) on the left titillator and 
4 to 7 teeth (usually 4, 5, or 6) on the right (Figs. 12-16). The teeth 
are closer together on the left titillator than they are on the right one 
(Figs. 12, 13). In another asymmetric touch, the left titillator is more 
dorsal than the right one (Fig. 12). Like the number of teeth, the length 
of the titillators varies individually instead of interspecifically or geo- 
graphically: in each species of the rhesus group, the titillators may be 
equal (Figs. 12, 13, 15), or the left one may be a little to a lot shorter 
than the right. Two of the three cornuti are conspicuous and tentlike— 
one essentially straight, the other slightly to very curved, each with 3 
to 6 teeth, which are closer together on the straight cornutus than they 
are on the curved one (Figs. 12-14). The third cornutus is very incon- 
spicuous, amounting to nothing but a bit of lightly sclerotized vesica 
(Fig. 12). 

Right down to the titillators and cornuti, the aedeagus of the rhesus 
group recalls that of the themistocles group (compare Figs. 12-16 with 
figs. 11-19 in MacNeill 1993 and see MacNeill’s diagnosis of the themis- 
tocles group). Some variation is less rampant in the themistocles group, 
even though that group is larger: in P. themistocles (Latreille) itself, 
the two titillators are always the same length (fig. 19 in MacNeill 1993); 
and in what I consider the peckius subgroup comprising P. peckius 
(Kirby), P. sabuleti (Boisduval), P. norae MacNeill, P. draco (Edwards), 
and P. mardon (Edwards), the left titillator is always much shorter 


36 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 15, 16. Male genitalia of rhesus group Polites (minus left valva) in left lateral 
view (ex Godman 1900:pl. 93, figs. 4, 7). 15, Polites subreticulata from MEXICO or 
GUATEMALA (aedeagus artificially rotated so that the left titillator is lower than the 
right whereas the reverse is really true); 16, Polites rhesus from Milpas, 5900 ft (1800 
m), Durango, MEXICO (tegumen plus uncus artificially twisted so that the underlying 
gnathos and the gap between it and the uncus fail to show). 


than the right (figs. 11-18 in MacNeill 1993). Throughout the themis- 
tocles group, the third, lightly sclerotized cornutus (inconspicuous in 
the rhesus group) is elongate and suggestive of a scouring pad (with a 
surface that MacNeill [1998] calls “minutely scobinate’’). Otherwise, 
the basic forms and arrangements of aedeagal accessories are closely 
similar in the rhesus and themistocles groups (small differences will 
be evident in the comparison of figures—but a few figures cannot 
adequately convey individual variation, which tends to weaken such 
differences). In contrast, forms and arrangements of aedeagal acces- 
sories differ sharply (and in different ways) in the origenes group (Figs. 
1-5), the vibex group, and P. baracoa. 

What aspects of the male genitalia set the rhesus group apart? In 
Polites the distal end of the uncus is divided: each of the two uncal 
tips terminates in a two-layered comb, with extremely close-set. tines 
curving backward and downward in each layer. These ““uncal combs” 
(called “‘pectines’” by MacNeill 1993) are slightly enlarged in the rhesus 
group (Figs. 11, 12, 14, 16), intermediate in the themistocles and ori- 
genes (Figs. 2-5) groups, reduced in the vibex group, and absent in P. 
baracoa. There is a sizable gap between the distal end of the uncus and 
the underlying divided gnathos in the rhesus (Figs. 12, 14, 15), origenes 
(Figs. 1, 8, 5), and vibex groups, and in P. themistocles (fig. 10 in 
MacNeill 1993)—but not in other members of the themistocles group 
(i.e., the peckius subgroup), where the gnathos is up against the bottom 
of the uncus (figs. 2-9 in MacNeill 1993). There is no gap in P. baracoa, 
either, but for a totally different reason: this species has lost the un- 
derlying divided gnathos. 

In Polites the dorsodistal corner of the valva is split by a small notch 
(Figs. 3, 5, 12, 14-16, and figs. 2-10 in MacNeill 1993): the dorsal 
margin of the valva immediately anterior to this notch supports a dense 
field of more or less dorsally-directed bristles; and the posterior margin 
of the valva immediately posterior to this notch displays (in lateral 
view) at least one sizable, dorsally- or dorsocaudally-directed tooth 


VOLUME 48, NUMBER 1 OT 


which, in turn, joins one or more teeth (readily visible in posterior view 
[Figs. 2, 4]) that extend mediad beneath the more or less mediaily- 
expanded posterior edge of the dense field of bristles. Medial expansion 
of the bristled area is least in the vibex and themistocles groups, strong 
in the origenes group, and greatest in P. baracoa and the rhesus group. 
In lateral view this expansion produces a slight hump (in the origenes 
group, Figs. 3, 5) to a very perceptible hump (in P. baracoa and the 
rhesus group, Figs. 12, 14) on the dorsal margin of the valva anterior 
to the valval notch. The posterior edge of the dense field of bristles is 
without teeth in the rhesus group, finely dentate in the themistocles 
group, more coarsely dentate in P. baracoa and the origenes group 
(Figs. 2, 4), and most coarsely dentate in the vibex group. 

About halfway down the posterior margin of the valva (in lateral 
view) a massive projection extends backward in P. baracoa and one or 
more sizable teeth point backward in the rhesus group (Figs. 12, 14- 
16). This portion of the posterior margin is finely dentate (Figs. 3, 5) 
to smooth in the origenes group and essentially smooth in the vibex 
and themistocles groups (figs. 2-10 in MacNeill 1998). 

The one or more teeth—visible in posterior view (Figs. 2, 4)—that 
extend mediad beneath the posterior edge of the dense field of bristles, 
are conspicuously multiple in the origenes group (Figs. 2, 4), in the 
vibex group, in P. baracoa, and in P. themistocles but are single to 
inconspicuously multiple especially in the peckius subgroup but also, 
somewhat less consistently, in the rhesus group. 

The posterior margin of the valva (in lateral view) has more of a 
“chin” —so looks more squared off—in the rhesus group (Figs. 12, 14- 
16) than in any other group of Polites. Certain members of the origenes 
group (P. mystic, P. sonora, P. pupillus [Fig. 5], and some individuals 
of P. origenes) come closest to this well-chinned look. Farthest from 
it, in a sense, is the vibex group because the posterior half of the ventral 
margin of the valva is narrowly excised. 

Female genitalia in the rhesus group (Figs. 17-22) are broadly rem- 
iniscent of those of the origenes group—particularly P. origenes, P. 
puxillius, and P. pupillus (Figs. 6-9)—except that the apophyses an- 
teriores are not sclerotically connected to the lamella postvaginalis. 
Sclerotization of the lamella postvaginalis is intermediate, as in P. or- 
igenes, P. puxillius, and P. pupillus. Sclerotization of the ductus bursae 
is partial—mostly ventral and lateral, with a narrow, dorsal extension 
anteriorly—in P. rhesus (Figs. 17, 18) and P. carus (Figs. 19, 20), much 
as in P. origenes, P. puxillius (Figs. 6, 7), and P. pupillus (Figs. 8, 9); 
but it is virtually complete in P. subreticulata (Figs. 21, 22). In all three 
species of the rhesus group, as in all members of the origenes group, 
the ductus bursae has a ventral, longitudinal groove or break. This 


38 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 17, 18. Female genitalia of Polites rhesus from Denver, Colorado, USA [X- 
3490] (USNM). Scale = 1.0 mm. 17, Ovipositor lobes (exserted) with apophyses posteriores, 
eighth tergite with apophyses anteriores (not sclerotically connected to the lamella post- 
vaginalis of the sterigma), sterigma, and bursa copulatrix in ventral view; 18, The same, 
plus part of the ductus seminalis, in right lateral view. 


groove is to the right of center in the rhesus group (Figs. 17, 19, 21) 
instead of more or less midventral as it is in the origenes group (Figs. 


6, 8). | 


Owing to their overall conservatism and their individual variation, 


VOLUME 48, NUMBER 1 39 


the male genitalia of the rhesus group are not diagnostic at the specific 
level. But here, as in the themistocles group, external color pattern 
separates species: the underside of the hindwing instantly sets P. rhesus 
(Figs. 27, 28) apart from P. carus (Figs. 29, 30) and P. subreticulata 
(Figs. 31, 32). These last two species are much more similar in pattern 
(so much so that they are mistakenly regarded as subspecies). Color 
may help in determining unworn specimens: the spots of the upperside 
are usually creamy to pale yellow in P. carus and light to medium 
yellow-orange in P. subreticulata. Although the undersides tend to be 
creamier in carus, yellower in subreticulata, they overlap considerably. 

It is the female genitalia that best distinguish these species. The ductus 
bursae is only about half sclerotized (ventrally and laterally) in P. carus 
(Figs. 19, 20) but almost fully sclerotized in P. subreticulata (Figs. 21, 
22). In both, the sclerotized ductus bursae usually looks rather globular 
in ventral view (Figs. 19, 21); but in P. subreticulata it tends—dorso- 
laterally—to extend backward slightly (and usually to flare slightly) at 
the ostium bursae (Fig. 21). Genitalic differences between P. carus and 
the superficially distinct P. rhesus are much more subtle because in 
rhesus, as in carus, the ductus bursae is only about half sclerotized 
(ventrally and laterally) (Figs. 17, 18). However, the sclerotized ductus 
bursae looks a little more elongate in P. rhesus, especially in ventral 
view (Fig. 17); and, where it approaches the corpus bursae, the anterior 
edge of the sclerotization is more irregular in rhesus (Fig. 18) than it 
is in P. carus (Fig. 20) (and, for that matter, P. swbreticulata [Fig. 22)). 

Polites rhesus ranges from southern Canada (Saskatchewan and AI- 
berta), in a fairly narrow strip through the western Great Plains and 
southern Rocky Mountains of the United States (Stanford & Opler 1993), 
to high mountains of central Mexico (10,000 ft [3050 m] in the state of 
Mexico); P. carus, from the southwestern United States (western Texas 
to southeastern California—see Stanford & Opler 1993) to central Mex- 
ico (Distrito Federal); and P. subreticulata, from central Mexico (Sina- 
loa, Jalisco, Colima, Michoacan, Distrito Federal, Morelos, and Vera- 
cruz), through Central America, to Panama (as far, at least, as the 
Canal). 


DISCUSSION 
Generic Limits and Vanishing Traits 


Extending generic limits this way calls for more discussion of variation 
in characters—especially their reduction and loss. The form of the 
antennal club and the length of its reflexed apiculus, which are widely 
used in skipper classification, are undeniably valuable. However, though 
they tend to be conservative at the generic level, they are hardly im- 


40 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 19, 20. Female genitalia of Polites carus from Sunny Glen Ranch, 5000-7000 
ft (1525-2135 m), near Alpine, Texas, USA, 1-15 May 1926 [X-3488] (USNM). Scale = 
1.0 mm. 19, Ovipositor lobes (exserted) with apophyses posteriores, eighth tergite with 
apophyses anteriores (not sclerotically connected to the lamella postvaginalis of the ste- 
rigma), sterigma, and bursa copulatrix in ventral view; 20, The same, plus part of the 
ductus seminalis, in right lateral view. 


VOLUME 48, NUMBER 1 Al 


Fics. 21,22. Female genitalia of Polites subreticulata from Mazatlan, Sinaloa, MEX- 
ICO, J. A. Kusche [X-3494] (USNM). Scale = 1.0 mm. 21, Ovipositor lobes (exserted) 
with apophyses posteriores, eighth tergite with apophyses anteriores (not sclerotically 
connected to the lamella postvaginalis of the sterigma), sterigma, and bursa copulatrix in 
ventral view; 22, The same, plus part of the ductus seminalis, in right lateral view. 


42 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 23-32. Adults of species moved to Polites (all x1) (in USNM unless otherwise 
indicated); in each figure, upperside on left, underside on right. 23, puxillius 6, 28 mi 
(45 km) E El Salto, 8400 ft (2560 m), Durango, MEXICO, 6-7 August 1972, Veirs & 
MacNeill (MacNeill collection); 24, puxillius 2, 25 mi (40 km) W Durango, 8100 ft (2470 
m), Durango, MEXICO, 20 July 1964, J. A. Chemsak [X-3196] (Univ. Calif. Berkeley 
collection); 25, pupillus 6, Sierra de Guerrero, MEXICO, July 1913, R. Miller [X-2699] 
(type of Amblyscirtes catahorma Dyar); 26, pupillus 2, 2 mi (3 km) SW Potrerillos, 4200 
ft (1280 m), Sinaloa, MEXICO, 7-8 August 1986, J. Brown & Powell [X-3203] (Univ. 
Calif. Berkeley collection); 27, rhesus 6, Sapillo Creek Valley, 5800 ft (1770 m), 12 mi 
(19 km) N Pinos Altos, Grant Co., New Mexico, USA, 28 May 1959, J. M. & S. N. Burns 
[X-2712]; 28, rhesus 2, Cedar Creek Canyon, 6900 ft (2100 m), Ruidoso, Lincoln Co., 
New Mexico, USA, 24 May 1959, J. M. & S. N. Burns [X-3504],; 29, carus 6, Portal, 
Chiricahua Mountains, 4800 ft (1465 m), Cochise Co., Arizona, USA, 18 July 1974, J. M. 
& S. N. Burns [X-2708]; 30, carus 2, 6.5 mi (10.5 km) NE Sawtooth Mountain, Davis 
Mountains, 5600 ft (1705 m), Jeff Davis Co., Texas, USA, 28 April 1959, J. M. & S. N. 
Burns; 31, subreticulata 6, Mexico City, MEXICO, 7 June 1897, O. W. Barrett [X-3498] 
(forewing with a long tear in space 1b); 32, swbreticulata 2, Mexico City, MEXICO [X- 
3491], 7 


VOLUME 48, NUMBER 1 43 


mutable. For example, Amblyscirtes alternata (Grote & Robinson), 
which is an Amblyscirtes in the narrowest sense, is unique among all 
those species in having the sizable, delicate, sharp apiculus suddenly 
reduced to a short, blunt fraction of itself (Burns 1990). Lumping 
Yvretta with Polites may bother those who overweigh antennal clubs 
because the very short but definite apiculus of the rest of Polites seems 
to be lacking in the rhesus group. (Its blunt antennal club is one of the 
main reasons why Yvretta was created in the first place and kept well 
removed from Polites.) But in most specimens of each of the species 
of the rhesus group the blunt club actually ends in a slightly reflexed 
nubbin (comprising a few segments) that looks like a variably vestigial 
apiculus. A supposedly important difference between the rhesus group 
and the rest of Polites becomes relatively trivial. 

Differences in stigmal expression are demonstrably insignificant. Al- 
though males of P. carus and P. subreticulata of the rhesus group have 
a fairly well developed, Polites-type stigma, males of P. rhesus do not: 
the stigma is, at best, variably vestigial, and is usually missing altogether. 
In apparent contrast, “‘real’’ Polites males “always” develop a good 
stigma—or they did until MacNeill (1993) described P. norae, which 
produces nothing at all. (Polites norae is near P. sabuleti in the stigmally 
well endowed themistocles group.) Stigmal expression has been shown 
to vary ina similar, unpredictable manner within other genera. Despite 
a respectable male stigma in most species of Atrytonopsis, the lunus 
group entirely lacks one (Burns 1982), and A. deva (Edwards) “runs a 
gamut from no stigma, through many and various vestigial and reduced 
expressions, all the way to the complete three-part structure’ (Burns 
1982:551). 

On the genitalic front, no importance can be attached to the fact 
that the uncal combs of the rhesus group are bigger than those of other 
Polites. They are only about as much larger than those of the themisto- 
cles and origenes groups as those of the vibex group are smaller. At the 
farthest extreme, Polites includes baracoa, whose uncal combs have 
vanished without a trace. Moreover, baracoa has lost its gnathos. And, 
in a different kind of unique development, its paired, toothed, caudally- 
extending titillators have broadly joined each other, near their anterior 
ends, across the underside of the aedeagus. Altogether, in its genitalic 
morphology, the rhesus group is considerably closer to the Polites main- 
stream than is P. baracoa. 

Some may be jolted by the broad, round, female-like wings of males 
of P. puxillius (Fig. 23) and P. pupillus (Fig. 25) within the origenes 
group, where males of all other species have narrower, more pointed 
wings than do females. But sexual dimorphism in wingshape (which is 
typical and extremely widespread in skippers) has abruptly disappeared 


44 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


elsewhere within the genus Polites—in P. mardon within the peckius 
subgroup of the themistocles group. 


Distribution and Speciation 


In the origenes group, P. origenes (mainly from the eastern and 
central United States) morphologically and geographically links the 
northern, transcontinental superspecies P. mystic (P. mystic and P. 
sonora) and the Mexican sister species P. pupillus and P. puxillius. 
These montane sisters appear to be closely allopatric, in part through 
different altitudinal preferences; but data are too few to say for sure 
(see Fig. 10). Phylogenetically closer to the Mexican sisters than to 
superspecies P. mystic, P. origenes is out of touch with them and broadly 
sympatric with the superspecies—mostly with P. mystic (see maps in 
Opler & Malikul 1992, Stanford & Opler 1993). However, across all 
five species of the group, sympatry is limited. Their spatial distribution 
strongly reflects a set of allopatric speciation events. 

In the rhesus group, the superficially distinctive and more northward 
ranging P. rhesus extensively overlaps P. carus in the southwestern 
United States and northern Mexico, whereas P. carus and P. subreticu- 
lata, which are sister species, seem (from meager distributional data) 
almost to replace each other geographically. 

Again, the four strictly western members of the peckius subgroup of 
the themistocles group—P. draco, P. sabuleti, P. norae, and P. mar- 
don—are essentially allopatric (MacNeill 1993). So are several members 
of the vibex group, which Evans (1955) erroneously treated as a single, 
very widely distributed polytypic species. 

All four species groups of Polites overlap in distribution, ranging in 
broadly repetitious—though different—patterns from the neotropics to 
the nearctic: the vibex group from Argentina, Paraguay, Brazil, Bolivia, 
and Peru to Mexico, the West Indies, and (primarily) the southeastern 
United States; the rhesus group from Panama to southern central Can- 
ada; the origenes group from southern Mexico to the width of southern 
Canada; and the themistocles group from central Mexico to the width 
of southern Canada, plus Yukon Territory. But all four species groups 
of Polites still give clear distributional evidence of allopatric speciation 
within themselves. This is independent evidence that the morpholog- 
ically defined species groups are correct. 


ACKNOWLEDGMENTS 


Frederick H. Rindge at the American Museum of Natural History, New York, New 
York, furnished some important material; Jerry A. Powell, Essig Museum of Entomology, 
University of California, Berkeley, John E. Rawlins, Section of Invertebrate Zoology, 
Carnegie Museum of Natural History, Pittsburgh, Pennsylvania, C. Don MacNeill, and 
Doug Mullins loaned some; and MacNeill, John Kemner, and H. A. Freeman donated 


VOLUME 48, NUMBER 1 45 


some. Don Harvey helped with the map (Fig. 10). Elizabeth Klafter and the late Adrienne 
Venables dissected genitalia (126 KOH-preparations of Polites, about evenly split between 
the sexes). Daniel Otte at The Academy of Natural Sciences, Philadelphia, Pennsylvania, 
made a digital image of one dissection (Fig. 14), Victor E. Krantz photographed adults 
(Figs. 23-32) as well as the Godman genitalic illustrations (Figs. 1, 15, 16), and Young 
Sohn drew eight dissections (Figs. 2-9, 11-18, 17-22) and then mounted all figures. Sarah 
Burns aided in many ways. George T. Austin and C. Don MacNeill perused the manuscript. 
My thanks to everyone. 


LITERATURE CITED 


BurRNS, J. M. 1964. Evolution in skipper butterflies of the genus Erynnis. Univ. Calif. 
Publ. Entomol. 37:1-217. 

1982. Lychnuchoides frappenda from central Mexico joins lunus and zweifeli 

in a lunus group of Atrytonopsis (Lepidoptera: Hesperiidae: Hesperiinae). Proc. 

Entomol. Soc. Wash. 84:547-567. 

1988. Superspecies Atrytonopsis ovinia (A. ovinia plus A. edwardsi) and the 

nonadaptive nature of interspecific genitalic differences (Lepidoptera: Hesperiidae). 

Proc. Entomol. Soc. Wash. 85:335-358. 

1990. Amblyscirtes: Problems with species, species groups, the limits of the 
genus, and genus groups beyond—A look at what is wrong with the skipper classi- 
fication of Evans (Hesperiidae). J. Lepid. Soc. 44:11-27. 

Evans, W. H. 1955. A catalogue of the American Hesperiidae indicating the classifi- 
cation and nomenclature adopted in the British Museum (Natural History). Part IV. 
Hesperiinae and Megathyminae. British Museum, London. 499 pp., pls. 54-88. 

GODMAN, F. D. 1900. In Godman, F. D. & O. Salvin. 1879-1901. Biologia Centrali- 
Americana; Insecta; Lepidoptera-Rhopalocera. Vol. 2, 782 pp.; Vol. 3, 113 pls. 

HEMMING, F. 1935. Notes on seventeen genera of Rhopalocera. Stylops 4:1-3. 

HOFFMANN, C. C. 1941. Catalogo sistematico y zoogeografico de los Lepidopteros 
Mexicanos. Segunda parte. Hesperioidea. Anales Inst. Biol. [Mexico] 12:237-294. 

LINDsEY, A. W., E. L. BELL & R. C. WILLIAMS JR. 1931. The Hesperioidea of North 
America. Denison Univ. Bull., J. Sci. Lab. 26:1-142. 

MACNEILL, C. D. 1993. Comments on the genus Polites, with the description of a new 
species of the themistocles group from Mexico (Hesperiidae: Hesperiinae). J. Lepid. 
Soc. 47:177-198. 

OPLER, P. A. & V. MALIKUL. 1992. A field guide to eastern butterflies. Houghton Mifflin 
Co., Boston, New York, London. xvii + 396 pp., 48 pls. 

SCUDDER, S. H. 1889. The butterflies of the eastern United States and Canada with 
special reference to New England. Publ. by the author, Cambridge, Massachusetts. 
Vol. 3, pp. vii + 1775-1958, pls. 1-89, 3 maps. 

SKINNER, H. & R. C. WILLIAMS JR. 1923. On the male genitalia of the Hesperiidae of 
North America. Paper III. Trans. Am. Entomol. Soc. 49:129-158. 

1924. On the male genitalia of the Hesperiidae of North America. Paper V. 
Trans. Am. Entomol. Soc. 50:141-156. 

STANFORD, R. E. & P. A. OPLER. 1993. Atlas of western USA butterflies, including 
adjacent parts of Canada and Mexico. Publ. by the authors, Denver and Fort Collins, 
Colorado. xi + 275 pp. 


Received for publication 6 August 1993; accepted 12 September 1998. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 46-50 


A NEW SPECIES OF ACROLEPIOPSIS AND THE 
DESCRIPTION OF THE FEMALE OF 
A. CALIFORNICA (ACROLEPIIDAE) 


REINHARD GAEDIKE 


Deutsches Entomologisches Institut (DEI), Schicklerstrasse 5, 
D 16202 Eberswalde, Germany 


ABSTRACT. The family Acrolepiidae, of the superfamily Yponomeutoidea, includes 
three genera: Digitivalua Gaedike, Acrolepiopsis Gaedike, and Acrolepia Curtis. Acro- 
lepiopsis liliitvora, new species, from California and Oregon, is described herein; the 
male and female genitalia are illustrated. Adults have been reared from the bulbs of 
Lilium washingtonianum. The female of A. californica, previously unknown, also is 
described, and the genitalia are illustrated. The larvae are reported to feed on Disporum 
hookeri. 


Additional key words: Acrolepiopsis liliivora, genitalia (male and female), Lilium 
washingtonianum, Disporum hookeri. 


Acrolepiidae is a family in the superfamily Yponomeutoidea. It is 
represented in all zoogeographical regions and includes 84 described 
species in three genera: Digitivalvua Gaedike (40 species), Acrolepiopsis 
Gaedike (35 species), and Acrolepia Curtis (9 species). In the Nearctic 
and Neotropical regions there are 15 described species, three in Digi- 
tivalva, ten in Acrolepiopsis, and two in Acrolepia (Gaedike 1984a, 
1984b). Undoubtedly, more detailed study of each zoogeographical 
region will increase the number of known species. 

The known life histories indicate that acrolepiid larvae are leaf- 
skeletonizers or miners of leaves, fruit, or bulbs of Asteraceae (Digiti- 
valva), Dioscoriaceae and Liliaceae (Acrolepiopsis), and Solanaceae 
(Acrolepia). While examining acrolepiid from the western United States, 
I discovered a new species of Acrolepiopsis and the previously unde- 
scribed female of A. californica. These are described below. 


Acrolepiopsis liliivora Gaedike, new species 


Wingspan 15-16 mm. Head dark brown, except vertex light brown; 
labial palpi dark brown. Thorax dark brown. Forewing dark brown, 
with white triangular spot, sometimes overlaid with dark scales, ex- 
tending obliquely forward from margin of dorsum at % distance from 
base, with minute white dot on dorsum at base of cilia, and with 
individual light scales scattered throughout the dark area; cilia beneath 
apex pale distally. Males somewhat lighter brown than females. Male 
genitalia (Fig. 1): saccus elongate, narrow, apically rounded; valva with 


— 


Fics. 1-4. Genitalia of Acrolepiopsis liliivora Gaedike, new species. 1, Male genitalia; 
2, Female genitalia (sterigma); 3, Female genitalia (signa); 4, Female genitalia (signa). 


AT 


VOLUME 48, NUMBER 1 


48 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


base broad, narrowest medially and expanded apically, costal margin 
concave; aedeagus more than 2.5 times valva length, broad basally, 
tapering to apex, with minute sclerotizations in vesica. Female genitalia 
(Figs. 2-4): Eighth segment with a pair of relatively narrow, somewhat 
clublike structures bearing setae on the broadly rounded base; ostium 
with rectangular sclerotization, the lower half more strongly sclerotized; 
ductus bursae strongly sclerotized over most of its length; corpus bursae 
with two long signa; signa slightly bent with dentate inner surface, the 
appearance variable depending upon the preparation. 

Type locality: California: Auto Rest. (I have been unable to determine 
the exact location.) 

Types: Holotype male: Auto Rest, Cal [ifornia] 18. 8.[19]18, on Lilium 
washingtonianum, Coll. David Griffiths (genital slide R. Gaedike No. 
2421). Paratypes: 1 male, 3 females, same data as holotype; 1 female 
Oregon, Santiam Natl. For., reared from bulb of Lilium washingto- 
nianum, emerged 9. X. 1931. The holotype and three paratypes are 
deposited in the U.S. National Museum of Natural History, Smithsonian 
Institution, Washington, D.C.; two paratypes are deposited in the 
Deutsches Entomologisches Institut (DEI) Eberswalde. 

Biology: Larva in bulb of Lilium washingtonianum (Liliaceae), pupa 
in a net-like cocoon. 

Remarks: All specimens are in poor condition. One female lacks 
forewings; another lacks a hindwing. The new species is closely related 
to A. californica. It differs in the coloration of the cilia below the apex, 
in having somewhat broader valvae and longer signa than californica. 


Acrolepiopsis californica Gaedike, 1984 
Entomol. Abh. Staatl. Mus. Tierk. Dresden 47(1983)10:183-184, Fig. 20. 


In material sent to me by J. Powell there were four specimens of this 
species, two of which were females. Because the female was previously 
unknown, I describe the female genitalia below. 

Female genitalia (Fig. 5): The pair of ventrolateral clublike structures 
bearing setae on the broadly rounded base, tapering to a point; ostium 
with a somewhat cup-shaped sclerotization, the lower half more strongly 
sclerotized; ductus bursae strongly sclerotized; signa short, with dentate 
inner surface. 

Material examined: Two males, two females, as follow: One male: 
Calif.: El Dorado Co., Blodgett Forest, 13 mi E Georgetown, 4000- 
4500’, 27/28.V.1978, leg. J. Powell. One male: Calif.: Siskyou Co., 
McCloud River at Ash Creek Rgr. Sta., 9.VI.1974, leg. J. Powell. One 
female: Calif.: Myers Flat, Humboldt Co., 14.VIII.1963, leg. J. Powell. 
One female: Calif.: Big Creek Reserve (UCNLWR), Monterey Co., 26./ 


VOLUME 48, NUMBER 1 49 


Fic. 5. Female genitalia of Acrolepiopsis californica. 


28.V.1987, leg. J. Powell. These specimens are the first records of this 
species since its description. 

Biology: J. Powell (in litt.) gives some remarks on the biology of this 
species: “... In late April [1990] I made additional collections of the 
larvae of Acrolepiopsis at Big Creek on Disporum hookeri and suc- 
ceeded in rearing a few adults. This confirms the hostplant for the 
species represented by one specimen that I sent you from 1987. The 
larvae sometimes start to eating the inflorescence but sometimes do not; 
later they skeletonize the leaves, feeding on either upper or lower 
surfaces.” 


50 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


ACKNOWLEDGMENTS 


I am much obliged to J. B. Heppner and an anonymous reviewer for their help with 
the manuscript. I thank Heppner for the loan of material from the Smithsonian Institution, 
Washington, D.C., and J. A. Powell for the loan of material from the University of 
California, Berkeley. 


LITERATURE CITED 


GAEDIKE, R. 1984a. Revision der nearktischen und neotropischen Acrolepiidae (Lepi- 
doptera). Ent. Abh. Staatl. Mus. Tierk. Dresden 47(1983)10:179-194. 

1984b. Die Epermeniidae und Acrolepiidae der danischen Sidamerika-Expe- 

ditionen 1978/79 und 1981/82 (Lepidoptera). Reichenbachia Staatl. Mus. Tierk. 

Dresden 22(16):125-181. 


Received for publication 17 December 1998; revised and accepted 27 July 1998. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 51-57 


A NEW SPECIES OF LAPARA (SPHINGIDAE) FROM 
SOUTHEASTERN UNITED STATES 


VERNON ANTOINE BROU JR. 
74320 Jack Loyd Road, Abita Springs, Louisiana 70420 


ABSTRACT. Lapara phaeobrachycerous, new species, is described and illustrated. 
The new species presently is known to occur in extreme eastern Louisiana and the state 
of Mississippi in the southeastern United States. The species can be distinguished from 
Lapara coniferarum (J. E. Smith) by its slightly smaller size, darker color, the presence 
of a single postcellular dash, narrower wings, shorter antennae, and different flight period. 


Additional key words: endemism, hawkmoths, Louisiana, voltinism. 


Hodges (1971) recorded two species of Lapara in North America 
north of Mexico: Lapara bombycoides Walker and Lapara coniferarum 
(J. E. Smith). He treated Lapara halicarnie (Strecker) as a synonym of 
L. coniferarum, and Lapara pinea Lintner as a synonym of L. bom- 
bycoides. In his review of Lapara, Riotte (1972) recognized L. halicarnie 
as distinct on the basis of features of the labial palpus, pretarsus struc- 
ture, color, size, maculation, genitalia, and larvae. Riotte also questioned 
the synonymy of L. pinea with L. bombycoides. Previous authors (e.g., 
Holland 1908, Clark 1919), also noted or discussed the problems asso- 
ciated with the “hypertrophied” type specimen of L. halicarnie. 

At the time of Riotte’s (1972) review, no Lapara species were known 
from west of the Mississippi River in the United States, and none was 
known to occur in Louisiana. Riotte examined 24] examples of L. 
coniferarum from 18 states, mostly bordering the east and Gulf coast, 
from New York and Rhode Island southward to Florida and westward 
to Mississippi. He also examined 649 examples of L. bombycoides from 
a much greater range, 48 examples of L. halicarnie from four south- 
eastern states, and the type of L. pinea from New York. 

Since 1972, 8422 specimens of Lapara from Louisiana have been 
collected by the author using ultraviolet light traps. Most of the better 
quality specimens are pinned, spread, and labeled, and are in the au- 
thor’s collection. From 1972 through 1985, 1946 specimens were col- 
lected and recorded as L. coniferarum. From 1986 through 1992, spec- 
imens were segregated into two phenotypes. During these seven years, 
6473 Lapara specimens were collected: 2247 (85%) are L. phaeobra- 
chycerous, new species, and 4226 (65%) are L. coniferarum. 


Lapara phaeobrachycerous Brou, new species 
(Figs. lA & B) 
Male. Head: Dark charcoal gray to occassional brownish gray in color, 
scales on front and vertex form a bluntly rounded area between an- 


o2 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fic. 1. The Lapara of Louisiana: L. phaeobrachycerous, A. male holotype, B. female 
allotype. L. coniferarum, C. male, D. female. Specimens pictured were collected at 
Louisiana, St. Tammany Parish, 6.8 km NE Abita Springs. 


tennae. Antennae laminate, biciliate, weakly hooked tip, length x = 
10.3 mm (9.0-12.0; n = 40). Thorax: Color above as described for head, 
below a slightly lighter shade. Forewing: Dorsal color charcoal gray, 
whitish scales mostly limited to area basad along curving postmedial 
line, usually one prominent postcellular dash, though a second dash to 
varying degrees of prominence is not uncommon. Ventral color usually 
lighter shade than above, unremarkable without bands or maculation, 
length x = 27.35 mm (25-81.5; n = 40). Hindwing: Unicolorous charcoal 
gray dorsally without whitish scales or maculation. Ventral surface as 
described for forewing. Genitalia: (n = 12) Valve generally oval, process 
of sacculus variable, narrow to broad projection, either acuminate, 
dentate, or combination of both, uncus apex mildly hooked (Fig. 2A). 

Female. Head: As described for male. Antennae simple, length x = 


VOLUME 48, NUMBER 1 53 


sew chi . 
is 
Sig | 


Fic. 2. Genitalia of L. phaeobrachycerous, Louisiana, St. Tammany Parish, A. male, 
B. female. 


10.6 mm (10.0-12.0; n = 18). Thorax: As described for male. Forewing: 
Dorsal and ventral color as described for male, length x = 31.6 mm 
(29.5-35.5; n = 14). Hindwing: Dorsal and ventral color as described 
for male. Genitalia: (n = 4) Sclerotized genital plate, rounded center 
convexity along distal edge. Entire structure unremarkable otherwise 
(Fig. 2B). 

Types. Holotype ¢ (Fig. 1A), USA, Louisiana, St. Tammany Parish, 
4.2 miles (6.8 km) NE Abita Springs, sec. 24T6SR12E, 9 Sept. 1991. 
Allotype ¢ (Fig. 1B), same locality, 19 Sept. 1991. Paratypes: 879 6 and 
18 2, same locality, April 1 to Oct. 28, 1983-92. Holotype and allotype 
deposited in U.S. National Museum of Natural History. Paratypes de- 
posited in Florida State Collection of Arthropods, Gainesville, Louisiana 
State University, Baton Rouge, and the author’s collection. 

Diagnosis. In contrast to L. phaeobrachycerous, scales on the front 
and vertex of the head of L. coniferarum form an acute distal projection 
between the antennae and lower margin of the frons, among Louisiana 
and Mississippi specimens. 

Maculation is variable within populations, especially among different 
broods. In Louisiana, individuals of the first brood of L. coniferarum 
are larger than those of subsequent broods, a trait seen in other Louisiana 
Sphingidae such as Isoparce cupressi (Bdv.). The first brood of L. coni- 
ferarum also exhibits the greatest variation in color and shade. Speci- 
mens can be very dark, or have a predominance of brown scales es- 
pecially on the forewings. It is not uncommon for darker specimens to 
exhibit a range of very dark gray to off-white or even near white 
hindwings. 


o4 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


In Louisiana, both Lapara species exhibit varying shades of gray or 
occasionally brownish gray. An area along the forewing inner margin 
between the transverse postmedial line and the base is chestnut brown 
in both species. In fresh specimens of coniferarum, the dorsal forewing 
ground color is usually ash gray. Unlike phaeobrachycerous, the entire 
dorsal surface of the forewing of coniferarum is suffused with white 
scales. In both species, these scales are especially concentrated basad of 
the along the postmedial line. The upper surface of the hindwing of 
coniferarum is noticeably lighter than the forewing, owing to white 
scales, which are more numerous near the base. This pattern occurs to 
a lesser degree in phaeobrachycerous but is absent in some specimens, 
the moths appearing unicolorous gray. Some coniferarum, usually in- 
dividuals of the first brood, also have unicolorous gray hindwings. 

Among the sparse forewing maculation of Larara are bold to faint 
black streaks occurring as a furcating transverse postmedial line, es- 
pecially on the veins between the postmedial line and the outer margin. 
These black vein lines flare out as they approach and include the fringe 
near each vein. The fringe between each vein is usually white in conifer- 
arum and white to off-white in phaeobrachycerous. 

The forewing maculation of both species includes a faint to bold 
median line, which arises from the chestnut-colored area along the 
inner margin and proceeds towards the apex. The upper one-third of 
this line abruptly turns inwardly and intersects the costal margin at 
varying angles, acutely to near perpendicular. In phaeobrachycerous, 
this line often intersects the costal margin more basad than seen in 
coniferarum. The sometimes faint antemedial line roughly parallels 
the median line. | 

Riotte (1972) noted that female L. coniferarum have less accentuated 
markings than males. Markings of both Louisiana species similarly 
exhibit less accentuated markings. The forewing postcellular dashes 
vary in number in both Louisiana species. In coniferarum there are 
usually two dashes, though one or three occur in varying degrees of 
prominence and with less frequency. Visually, males of the two species 
are easy to distinguish. Male phaeobrachycerous appear as darker, slightly 
smaller, short and narrow-winged specimens with short antennae. The 
antennae of male coniferarum in Louisiana average 29% longer, x = 
13.3 mm (11.5-14.5; n = 40), than in phaeobrachycerous. Differences 
in both wing length and antennae length of coniferarum and phaeo- 
brachycerous are statistically significant as defined by t-test (highest 
level a = 0.0005). In males of coniferarum, the antennae cilia are twice 
the length of those on phaeobrachycerous, while the shaft diameter is 
roughly equivalent in both species. 

In Louisiana, the average forewing lengths for coniferauam are: 


Ol 
Or 


VOLUME 48, NUMBER 1 


males x = 29.25 mm (27.5-31.5; n = 40), females x = 31.1 mm (29.5- 
35.0; n = 23). Riotte (1972) listed measurements for coniferarum as 
males 24-30 mm (n = 40), females 27-36 mm (n = 8), with no averages 
given. For L. halicarnie, Riotte listed forewing lengths as males 29-36 
mm (n = 18), females 32-40 mm (n = 6). 

The genitalia of L. coniferarum have been illustrated by Rothschild 
and Jordan (1903), Hodges (1971), and Riotte (1972). Both Riotte and 
Hodges discuss the variability of certain structures of the male genitalia, 
and Riotte illustrates the variation in the sacculus. Comparing female 
specimens from Louisiana, the distal edge of the lamella postvaginalis 
of L. coniferarum is gently rounded, without the degree of convex 
protrusion noted in L. phaeobrachycerous. This genital plate attribute 
does vary slightly over L. coniferarum’s vast range. 

Distribution. Despite extensive collecting throughout Louisiana, L. 
coniferarum has been taken only in six upper, southeastern parishes, 
all east of the Mississippi River: West Feliciana, East Feliciana, East 
Baton Rouge, Ascension, Tangipahoa, and St. Tammany. This region 
of the state is known as the Florida Parishes. : 

I have studied hundreds of Lapara specimens from Mississippi to 
Florida, and upwards along the east coast states to Pennsylvania. From 
Georgia and South Carolina, specimens exist along with coniferarum 
which appear assignable to neither coniferarum nor bombycoides. They 
appear as small, narrow-winged specimens, like phaeobrachycerous, 
and may account for prior literature records indicating bombycoides 
occurring south to Florida. Due to limited material, it is unclear if these 
specimens have a relationship to phaeobrachycerous, though I suspect 
they are indeed phaeobrachycerous. All specimens examined from the 
area around Clemson, South Carolina, were melanic (very dark charcoal 
gray to near black) narrow-winged small specimens with short antennae. 

The verified range of L. phaeobrachycerous includes only the states 
of Louisiana and Mississippi at present. Besides the type locality, spec- 
imens have been collected at Fluker, Tangipahoa Parish, Louisiana, 
and from Harrison, Pike, and Lee counties in Mississippi. 

Etymology. The specific epithet is derived from the Greek phaeo, 
meaning dark, dusky, and brachycerous, meaning short horns or an- 
tennae. 


DISCUSSION 


Lapara is a difficult genus for several reasons. Most field collected 
specimens are worn because the moths have a tendency to flutter along 
the ground through brush and leaf litter for considerable distances, 
especially when they approach light traps operating within a few feet 
of ground level. This problem appears to be reduced by operating traps 


06 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


at a greater height above ground, for example, above five meters. 
Females comprised less than one percent of the total Lapara specimens 
collected. Apparently, females are not attracted to ultraviolet light to 
the degree males are. 

In Louisiana, flight periods of L. coniferarum are different from 
those of L. phaeobrachycerous, though both species have five annual 
broods. Based on dates of capture over a seven-year period (1986-92), 
L. coniferarum has its initial annual brood peaking mid-April, with 
specimens beginning to appear in mid-March, and on occasion even as 
early as late February. The initial brood peak is separated from the 
second brood peak by an approximately two-month interval. Brood 
peaks two through four begin around mid-June and occur at 30-day 
intervals, In contrast, L. phaeobrachycerous has its first brood peak 
about mid-May, though the numbers of individuals of this brood are 
usually quite low compared to subsequent broods. All five broods occur 
at 30-day intervals. 

Koebele (1881) stated that L. coniferarum has at least two annual 
broods in the southern United States. Riotte (1972) also stated conife- 
rarum has two distinct flight periods in the southeastern coastal states. 
Those authors apparently did not recognize that their second brood/ 
flight period involved multiple sequentially occurring broods. 

Based on specimens collected at ultraviolet light traps, the nightly 
flight time for male coniferarum in Louisiana begins about midnight 
and lasts for approximately three to four hours, while females have 
appeared only at dusk. No flight time information is known about 
phaeobrachycerous. : 

Nothing is known about the immature stages of phaeobrachycerous, 
but I suspect that the larvae may feed on Pinus species. The area in 
which this new species occurs in Louisiana is classified as a longleaf 
pine region. 

Specimens fade quite rapidly. Pinned specimens several years old 
appear brown, quite different in color than freshly collected specimens. 
This is due in part to ultraviolet degradation as well as the effects of 
chlorine gas from vaporizing insect pest fumigants in storage cases (Brou 
1991). Descriptions of colors and other attributes are representative of 
those occurring on freshly collected and dried Louisiana specimens, 
unless stated otherwise. Clark (1919) commented that the type of L. 
halicarnie in Strecker’s collection appears faded. 


ACKNOWLEDGMENTS 


I thank the following individuals who supplied specimens, records, or aided in this 
project: Rick Kergosien, the late Charles Kimball, Bryant Mather, Richard S. Peigler, Eric 
L. Quinter, and J. C. E. Riotte. 


VOLUME 48, NUMBER 1 o7 


LITERATURE CITED 


Brou, V. A. 1991. Health related information on chemicals commonly used by ento- 
mologists and adjunct methods of pest control in stored entomological collections. 
South. Lepid. News 13:1-3. 

CLARK, B. P. 1919. Some undescribed Sphingidae. Proc. New England Zool. Cl. 6: 
67-68. 

Hopces, R. W. 1971. The moths of America north of Mexico, Fasc. 21 Sphingoidea. 
E. W. Classey Ltd. and R.B.D. Publications. 158 pp. 

HopcEs, R. W. et al. 1983. Check list of the Lepidoptera of America north of Mexico. 
E. W. Classey Ltd. and the Wedge Entomological Research Foundation, London. 
xxiv + 284 pp. 

HOLLAND, W. J. 1903. The moth book. Doubleday, Page and Co., New York. xxiv + 
479 pp. 48 plates. 

KOEBELE, A. 1881. Descriptions of and notes upon various larvae. Bull. Brooklyn En- 
tomol. Soc. 4:20-22. 

RIOTTE, J. C. E. 1972. A review of the North American hawk moth genus Lapara 
(Lepidoptera: Sphingidae). Life Sci. Contr., Royal Ont. Mus. No. 79. 40 pp. 


Received for publication 10 December 1992; revised and accepted 18 September 1993. 


PROFILE 


Journal of the Lepidopterists’ Society 
48(1), 1994, 58-67 


WILLIAM WITTFELD: THE FLORIDA CONNECTION 


JOHN V. CALHOUN! 
1731 San Mateo Drive, Dunedin, Florida 34698 


ABSTRACT. William Wittfeld (1828-1913) collected numerous natural history spec- 
imens, including Lepidoptera, at “Indian River, Florida,” in the vicinity of Georgiana, 
Brevard County. Many of his Lepidoptera specimens were sent to William H. Edwards, 
Henry Edwards, and Berthold Neumoegen. No fewer than thirty-three taxa of Lepidop- 
tera were described from material collected by Wittfeld. He contributed to the life history 
descriptions of at least nineteen species. Three patronyms of Lepidoptera honor Wittfeld. 
His daughter, Annie M. Wittfeld (1865-1887), also reared numerous species. 


Additional key words: Lepidoptera, “Indian River, Florida,” type locality, life his- 
tory, Annie M. Wittfeld. 


During the late nineteenth century, Florida was still largely a vast, 
undeveloped frontier. New railroad systems were allowing pioneers to 
establish homesteads in formerly remote Indian territory. In 1880, the 
entire population of Florida totalled less than 270,000 (Tebeau 1980). 
In March of that year, William Wittfeld (1828-1913) began collecting 
Lepidoptera at his home on “Indian River, Florida.” Wittfeld’s spec- 
imens and observations of Florida species became invaluable to the 
prominent Lepidopterists of the period. For more than a decade, “In- 
dian River, Florida” was a renowned source of unique and undescribed 
species of Lepidoptera. 

William Wittfeld (Fig. 1) was born in Germany on 3 April 1828. In 
1853, when he was 25 years old, Wittfeld immigrated to the United 
States, joining thousands of others seeking to escape political unrest in 
Germany. He left behind a large family, including sisters Hermine and 
Marie and brothers Herrmann, Hoeinreich, and Peter (Peter immi- 
grated to the United States in 1869). William lived in Philadelphia 
where he worked as a bookkeeper. Here, he married Rosalie Gottlieb 
in 1863, with whom he had two children: Annie, born in 1865, and 
Harry, born in 1867. During Wittfeld’s residency in Philadelphia, the 
city experienced “the coldest winter on record” (Weigley 1982). From 
that time forward, Wittfeld probably yearned for a milder climate. 

Lured by the prospects of comfortable temperatures and available 
land, the Wittfelds moved in 1869 to Merritt Island, Florida where they 


' Research Associate, Florida State Collection of Arthropods, Florida Department of Agriculture and Consumer Services, 
Gainesville, Florida. 


VOLUME 48, NUMBER 1 09 


ints Hae ; sv ne © 5 ae .* Be. Sd mn a fis 
Fic. 1. William and Rosalie Wittfeld at “Fairyland” (ca. 1890). 


were among the first settlers (Hellier 1965). In May 1870, Wittfeld 
applied for a federal land patent to secure 376 hectares (152 acres) of 
fertile land on what is now the southern end of Merritt Island, just south 
of the town of Georgiana in Brevard County. The property rose in a 
long slope from the Indian River (an inshore marine habitat, now part 
of the Intracoastal Waterway), eastward across a narrow peninsula to 
the shores of the Banana River (a marine lagoon). Wittfeld could view 
both bodies of water from the top of this slope, which he called Hon- 
eymoon Hill. Into the side of the hill, Wittfeld dug a hurricane shelter 
nearly large enough to stand in. Forests of live oak (Quercus virginiana 
Mill.) and cabbage palm (Sabal palmetto (Walt.) Lodd ex Schultes) 
dominated the hill overlooking a clear lake that Wittfeld appropriately 
named Honeymoon Lake. 

Wittfeld cultivated tropical fruits such as pineapples, bananas, and 
citrus. He also grew a variety of tropical plants that he generously 
distributed to his neighbors. His home was built next to a large Indian 
midden and burial mound and for many years was considered a “‘show- 
place” on the island (McAleenan 1991). Wittfeld started a school for 
his children next door to their home, bringing in a teacher who lived 
in a room adjoining the classroom. Over the years, Wittfeld developed 
his property into what he called “Fairyland” (or “Fairy Land’’), which 


60 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


2 \ 
W Merritt Island 


~f Wi Rockledge 


Banana River 


"Fairyland" 


Je 


Fic. 2. Location of “Fairyland” in Brevard County, Florida. 


became a popular tourist attraction during the late 1880’s (Fig. 2). 
Tourists rented rooms within five cottages built adjacent to Wittfeld’s 
home and strolled along trails that meandered through the local forest 
and around Honeymoon Lake. Although Wittfeld preserved much of 
the original landscape, developmental pressures began to have an effect 
on surrounding properties as early as the 1890’s. Holland (1898) and 
Skinner (1907) both mentioned the destruction of a natural area near 
Wittfeld’s home due to expanding cultivation. 

At least seven different spellings of Wittfeld’s name have been em- 
ployed. William H. Edwards consistently misspelled his name as ““Wiss- 
feld’’ in letters to Henry Edwards during 1880 and 1881. This spelling 
was published at least three times (W. H. Edwards 1881a, 1881b, Neu- 
moegen 1881) and was the result of Wittfeld’s own signature which 
closely resembled “‘Wissfeld.”” Additional spelling variations include 
Whitfeld (McAleenan 1991), Whitfeldt (Langlais 1984), Whittfeld 
(Anonymous 1918), Whitfield (Kjerulff 1972), Wittfield, and Whittfield 
(W. H. Edwards 1882d, H. Edwards 1883b, U.S. Census Bureau 1900, 
Hellier 1965). Although he was usually referred to as “Dr. Wittfeld” 
and W. H. Edwards (1881f) stated that Wittfeld was a “physician,” it 
is unclear whether he ever practiced medicine or the title was granted 
out of respect for his horticultural expertise (e.g., “herb doctor’’). Witt- 
feld was congenial, educated, spoke eloquent English, and had a pre- 
dilection for smoking cigars. 


VOLUME 48, NUMBER 1 61 


Wittfeld’s botanical interests eventually led to a fascination with 
insects, especially Lepidoptera. In March 1880, Wittfeld contacted W. 
H. Edwards about his desire to collect insects. Edwards persuaded his 
sister and her husband, who resided at Lake Jessop, Florida, to visit the 
Wittfelds during a trip to the Indian River region. She convinced Witt- 
feld to collect Lepidoptera specimens for Edwards who generously 
forwarded the necessary equipment including nets, poison, and insect 
pins. It appears that Wittfeld also agreed to collect a limited number 
of Coleoptera specimens at the request of W. H. Edwards, but the 
disposition of these specimens is unknown. 

Shipments of specimens from Wittfeld were eagerly accepted by 
Edwards who retained only the butterflies, offering the moths (and 
duplicate butterflies) to Henry Edwards and Berthold Neumoegen. As 
an incentive to Wittfeld, W. H. Edwards suggested that H. Edwards 
and Neumoegen begin contributing money to Wittfeld for the speci- 
mens they received (“it will be well and encouraging’’). To facilitate 
this endeavor, W. H. Edwards proposed a fixed price of 10 cents each 
for the desirable specimens. Edwards collected these payments and 
forwarded them directly to Wittfeld. Over the next twelve years, Witt- 
feld and W. H. Edwards developed a close relationship. 

During 1880 and 1881, Wittfeld supplied thousands of specimens to 
W. H. Edwards, H. Edwards, and Neumoegen. Individual shipments 
consisted of 500 or more specimens. In a letter dated 10 December 
1880, W. H. Edwards proposed to H. Edwards that Wittfeld could 
supply them four thousand specimens during 1881 for a fee of two 
hundred dollars. W. H. Edwards continued to instruct Wittfeld in 
proper collecting techniques and sent additional equipment especially 
suitable for collecting moths. 

As with many nineteenth century naturalists, Wittfeld’s passion to 
collect was not limited to one group of organisms. In February 1881, 
Wittfeld contacted Spencer F. Baird of the Smithsonian Institution and 
offered to collect snake specimens for the National Museum. He ex- 
plained that collecting Lepidoptera and Coleoptera “does not occupy 
all my time.” Wittfeld ultimately supplied Baird with numbers of small 
mammals, snakes, turtles, frogs, lizards, fish, shells, and a few insects 
(Coleoptera and Orthoptera; to be forwarded to W. H. Edwards if 
unwanted). All these specimens were shipped in jars of alcohol which 
Baird supplied. Wittfeld also procured some Indian artifacts for the 
museum. Baird paid Wittfeld for this material and based compensation 
on rarity. 

It is apparent that Wittfeld’s inexhaustible collecting activities oc- 
casionally exceeded the demand for his specimens, especially of ““com- 
mon” species. In late 1881, Henry Edwards complained about having 


62 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


to pay Wittfeld for all the specimens he had received. In response, W. 
H. Edwards wrote Wittfeld and asked him to reduce the number of 
shipments and to “collect only the rarest in such as he conveniently 
can and not give all his time to it as he has been doing.” S. F. Baird 
a'so advised Wittfeld (letter dated 27 March 1882) that many of the 
specimens he was sending were of little value to the Smithsonian col- 
lections. Despite the consistency of Wittfeld’s shipments, payments were 
sometimes painfully tardy. Delinquent payments were further exacer- 
bated by uncontrollable postal delays. W. H. Edwards lamented that 
Wittfeld’s letters were “sometimes 10 or 11 days in getting to me... 
and mine to him just the same.” Hardships associated with life in Florida 
also contributed to delays in shipments and payments. 

In 1881, Wittfeld bemoaned the hordes of mosquitoes he encountered 
whenever he attempted to collect, resulting in a reduction in the number 
of specimens he was able to secure. It is notable that one of the highest 
densities of mosquitoes ever recorded in Florida was near Wittfeld’s 
home in Brevard County where early residents of Merritt Island sup- 
posedly escaped mosquito bites by stuffing their clothing with crumpled 
newspaper (Myers and Ewel 1990). In September 1880, the Wittfelds 
were nearly devastated by a hurricane. W. H. Edwards was greatly 
concerned about their welfare and wrote (letter to H. Edwards dated 
11 September 1880) that he feared that Wittfeld “suffered by the late 
cyclone on that coast. Since that occurred I have had no letter from 
him. He has not missed a mail for months.’ Edwards’ fears were par- 
tially realized when Wittfeld confirmed that the hurricane “nearly 
ruined them.” The damage was severe. The storm “blew all the leaves 
off trees, tore down fences and buildings (but not his house), broke off 
all ripe oranges, broke flat the bananas” and “pulled pine apples from 
the roots.”’ As a result of the hurricane, Wittfeld temporarily suspended 
shipments of specimens to Edwards. The storm had damaged the mail 
boat, causing the mail to run so irregularly that he could not trust boxes 
to it. Surprisingly, Wittfeld resumed shipments to Edwards by early 
October of that year. 

Wittfeld soon became concerned that the increasing popularity of 
his captures would result in unwanted requests for specimens. In April 
1880, he asked W. H. Edwards to keep his name as confidential as 
possible. Only W. H. Edwards, H. Edwards, Neumoegen, and E. T. 
Cresson, Sr. were initially aware of Wittfeld’s identity. In January 1881, 
W. H. Edwards reminded H. Edwards not to publish Wittfeld’s name 
“for fear that it will lead to everybody writing him for butterflies.” 
Four months later, W. H. Edwards inexplicably ignored his own advice 
and published Wittfeld’s name (as ““Wissfeld’’) (W. H. Edwards 1881a). 

In early 1882, Wittfeld sought to provide specimens of Papilio to the 


VOLUME 48, NUMBER 1 63 


German entomologist, Otto Staudinger, while reserving the right to 
continue supplying material to W. H. Edwards and Neumoegen. After 
1888, shipments from Wittfeld became increasingly irregular. Very few 
shipments were sent during the late 1880’s. The correspondence be- 
tween Wittfeld and W. H. Edwards appears to have ceased in 1892. 
The Smithsonian Institution continued to receive small mammals and 
reptiles from Wittfeld until at least 1894. 

No fewer than 383 taxa of Lepidoptera were described from Wittfeld 
material and possess type localities of “Indian River, Florida.’ They 
are currently included in the families Tineidae (Beutenmueller 1887), 
Yponomeutidae (H. Edwards 1881la), Sessiidae (H. Edwards 1883b), 
Hesperiidae (W. H. Edwards 1880a, 188le, Lintner 1881, Skinner 1896), 
Pieridae (Skinner 1894, Miller and Brown 1981), Lycaenidae (W. H. 
Edwards 1883), Nymphalidae (W. H. Edwards 1880b), Limacodidae 
(H. Edwards 1886), Pyralidae (Hulst 1886, Kimball 1965), Geometridae 
(Grote 1882, Kimball 1965), Arctiidae (H. Edwards 1882, 1883a, Neu- 
moegen 1881), Apatelodidae (H. Edwards 1886), Sphingidae (Butler 
1881), and Noctuidae (Grote 1884, H. Edwards 1881b, 1886, Kimball 
1965). Wittfeld also discovered a new species of muskrat (Rodentia: 
Cricetidae) (True 1884). 

Wittfeld was the first to confirm the occurrence of several species of 
Lepidoptera in North America, including Battus polydamas (L.) (W. 
H. Edwards 1882a, 1882d), Hypolimnus misippus (L.) (W. H. Edwards 
1881d, 1888b), and Siproeta stelenes (L.) (W. H. Edwards 1885). Ed- 
wards was constantly impressed by the species that Wittfeld found, 
telling H. Edwards (letter dated 17 July 1881) ‘“‘one Cuban or Mexican 
species after another turns up there.” In addition to supplying dried 
adult specimens, Wittfeld also reared many species and sent W. H. 
Edwards live and preserved examples of ova, larvae, and pupae. Many 
of the preserved early stages of moths were forwarded to H. Edwards. 

W. H. Edwards personally reared many species of Lepidoptera from 
ova and larvae supplied by Wittfeld. Edwards had a particular fondness 
for Heliconius charitonius (L.). With the assistance of Wittfeld, Ed- 
wards reared this species numerous times and published a description 
of its early stages (W. H. Edwards 1881c). On 27 May 1880, Edwards 
wrote “I turned a 6 Heliconia Charitonia loose (from chrysalis) in my 
parlor yesterday and it was pretty to see it course about just as I 
remember adult Heliconias on the Amazon.” The behaviors of adult 
H. charitonius were carefully documented by Wittfeld and published 
by W. H. Edwards (1881f). Wittfeld’s specimens and detailed obser- 
vations contributed to the life history descriptions of at least nineteen 
species of Lepidoptera (see W. H. Edwards 188 1a, 1881b, 1882b, 1882c, 
1887-97, H. Edwards 1887, Scudder 1889). W. H. Edwards (1888) 


64 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


wrote that Wittfeld “has done more to elucidate the biological history 
of the Lepidoptera of Florida than any one who has preceded him.” 

Additional references to Wittfeld, “Indian River’ or Georgiana, Flor- 
ida can be gleaned from over one hundred years of literature on Lep- 
idoptera. Such sources include W. H. Edwards (1881g, 1884b), French 
(1885), Maynard (1891), Skinner (1911, 1921), Grossbeck (1917), Skin- 
ner and Williams (1924, 1925), Burns (1964) and Kimball (1965). Hol- 
land (1898, 1908, 1931) photographed several of Wittfeld’s specimens 
for his plates. Wittfeld’s specimens are also discussed in detail by Brown 
(1967, 1970), Brown and Miller (1975, 1977, 1980, 1987), and Calhoun 
(1993). Most major North American institutional collections possess 
Wittfeld Lepidoptera specimens. 

Annie M. Wittfeld shared her father’s interest in Lepidoptera. In 
September 1884, Annie documented the strange effects that a lightning 
strike had on the larvae of Limenitis archippus floridensis Strecker (W. 
H. Edwards 1884a). She was a talented artist and occasionally supplied 
drawings of new captures and early stages of Lepidoptera to W H. 
Edwards. Some of the life history information attributed to William 
Wittfeld is actually referable to Annie. In November 1887, Annie ob- 
served H. misippus ovipositing on purslane (Portulaca sp.) and sub- 
sequently reared the species on this plant (W. H. Edwards 1888b, dos 
Passos 1951). This remains one of only two known records of H. mis- 
ippus reproducing in North America. W. H. Edwards (1888b) char- 
acterized Annie as “intelligent and painstaking.’ Unfortunately, on 10 
April 1887, Annie died of rheumatic fever at the age of 23 (W. H. 
Edwards 1888a). Fifteen months earlier, on 10 January 1887, the Witt- 
felds’ only son, Harry, died suddenly of meningitis at the age of 20 
after becoming ill on a boat trip down the Indian River. William and 
Rosalie were devastated by these losses. It is likely that these tragedies 
contributed to William’s decreasing contact with W. H. Edwards and 
the Smithsonian Institution during the late 1880's. 

The final blow to William occurred with the death of his wife on 16 
December 1906. She was interred with Annie and Harry under a large 
oak tree near their home at “Fairyland.” In 1911, their remains were 
relocated to the Georgiana Cemetery. William’s health gradually failed. 
During the closing years of his life, he was cared for at the home of 
his friends, Mr. and Mrs. John Frye, of Rockledge, Florida. It was here 
that William Wittfeld died of edema on 13 July 1918 at the age of 86. 
We was interred beside his wife and two children at Georgiana Cem- 
etery. 

Today, there is little evidence of “Fairyland” or the habitats that 
Wittfeld frequented. Honeymoon Lake is nearly surrounded by housing 


VOLUME 48, NUMBER | 65 


developments. Nevertheless, neighboring properties still support trop- 
ical plants that grew from Wittfeld’s cuttings. 


Patronyms in Lepidoptera named in honor of William Wittfeld: 


wittfeldii, W. H. Edwards, Thecla, 1883, Canadian Entomol. 15:136-137. 
wittfeldii, H. Edwards, Alypia, 1883, Papilio 3:34. 
wittfeldii, H. Edwards, Pyrohotaenia, 1883, Papilio 3:156. 


ACKNOWLEDGMENTS 


I thank the late Theresa Bier, Nancy C. Sieck, Lucy Tomdale, Bill Wittfeldt, Rose 
Wooley, and Vera Zimmerman. Without their invaluable assistance, this biography could 
not have been possible. Lucy Tomdale generously supplied the photograph of William 
and Rosalie Wittfeld. Lee D. Miller and Jacquiline Y. Miller of the Allyn Museum of 
Entomology, Florida Museum of Natural History, kindly granted me access to copies of 
letters written by William H. Edwards. The West Virginia State Archives supplied copies 
of the two remaining Wittfeld letters in the W. H. Edwards files. Mark Epstein and 
William Cox provided copies of correspondence between Wittfeld and officials of the 
Smithsonian Institution which are deposited in the Smithsonian Institution Archives. I 
also express thanks to Alice Sanders, Librarian at the Division of Plant Industry, Florida 
Department of Agriculture and Consumer Services, who patiently helped me obtain 
numerous literature references. Finally, thanks are extended to Marc Epstein and William 
D. Winter for critically reviewing the manuscript. 


LITERATURE CITED 


ANONYMOUS. 1913. [Obituary of Dr. William Wittfeld]. East Coast Advocate (8 
Aug.):5. 

BEUTENMUELLER, W. 1887. Descriptions of new species of North American Tineidae. 
Entomol. Am. 3:139-140. 

BROWN, F. M. 1967. The types of the nymphalid butterflies described by William Henry 
Edwards—Part III, Nymphalinae, Limenitidinae, Apaturinae and Charaxinae. Trans. 
Am. Entomol. Soc. 93:319-393. 

1970. The types of the lycaenid butterflies described by William Henry Edwards. 
Part II—Theclinae and Strymoninae. With four neotypes established by Paul A. 
Opler. Trans. Am. Entomol. Soc. 96:19-77. 

Brown, F. M. & L. D. MILLER. 1975. The types of the hesperiid butterflies named by 
William Henry Edwards. Part I, Hesperiidae; Pyrginae. Trans. Am. Entomol. Soc. 
101:597-649. 

1977. The types of the hesperiid butterflies named by William Henry Edwards. 

Part II, Hesperiidae: Hesperiinae, Section I. Trans. Am. Entomol. Soc. 103:259-302. 

1980. The types of the hesperiid butterflies named by William Henry Edwards 

Part II, Hesperiidae: Hesperiinae, Section IJ. Trans. Am. Entomol. Soc. 106:43-88. 

1987. The types of the hesperiid butterflies described by William Henry Ed- 
wards. Part II—Hesperiidae: Hesperiinae, Section III and Megathymidae. Trans. Am. 
Entomol. Soc. 113:29-71. 

Burns, J. M. 1964. Evolution in skipper butterflies of the genus Erynnis. Univ. Calif. 
Publ. Entomol. 37:1-216. 

BUTLER, A. G. 1881. Notes on some North American Lepidoptera. Papilio 1:103-106. 

CALHOUN, J. V. 1993. Designation of a lectotype of Nisoniades somnus and notes on 
the occurrence of Erynnis icelus in Florida (Hesperiidae). J. Lepid. Soc. 47:49-54. 

DOs Passos, C. F. 1951. The entomological reminiscences of William Henry Edwards. 
J. New York Entomol. Soc. 59:129-186. 

EDWARDS, H. 188la. A new genus and some new forms of North American Zygaenidae. 
Papilio 1:80-81. 


66 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


1881b. Descriptions of some new species of Heterocera. Papilio 1:115-121. 
1882. New specie of Heterocera. Papilio 2:9-15. 

1883a. New forms of the genus Alypia. Papilio 3: 33-34. 

1883b. New species of Aegeriadae. Papilio 3:155-157. 

1886. Notes on North American Zygaenidae and Bombycidae with descriptions 
of new forms. Entomol. Am. 2:8-15. 

1887. Early stages of some North American Lepidoptera. Entomol. Am. 3:161- 


Wl 

EDWARDS, W.H. 1880a. Description of a new species of Pamphila from Florida. Canad. 
Entomol. 12:224-225. 

1880b. Description of a new species of Limenitis. Canad. Entomol. 12:246- 


291. 


188la. Description of the preparatory stages of Apatura flora, Edw. Canad. 

Entomol. 13:81-85. 

1881b. Description of the preparatory stages of Papilio palamedes, Drury. 

(Calchas, Fab.). Canad. Entomol. 13:119-123. 

1881lc. Description of preparatory stages of Heliconia Charitonia, Linn. Canad. 

Entomol. 13:158-162. 

1881d. Capture of Diadema bolina, Linn, in Florida. Papilio 1:30. 

188le. Description of a new hesperian from Florida. Papilio 1:78-79. 

1881f. On certain habits of Heliconia Charitonia, Linn., a species of butterfly 

found in Florida. Papilio 1:209-215. 

1881g. Notes on the species of Callidryas found within the United States. Trans. 

Am. Entomol. Soc. 9:9-14. 

1882a. Note on Papilio polydamas, Linn. Canad. Entomol. 14:120. 

1882b. Description of the preparatory stages of Neonympa areolatus, Smith- 

Abbot. Canad. Entomol. 14:163-166. 

1882c. Description of the preparatory stages of Grapta interrogationis, Fab. 

Canad. Entomol. 14:201-207. 

1882d. Papilio polydamas in Florida. Papilio 2:122. 

1883. Description of a new species of Thecla from Florida. Canad. Entomol. 

15:1386-137. 

1884a. [Letter from Annie M. Wittfeld]. Canad. Entomol. 16:180. _ 

1884b. Revised catalogue of the diurnal Lepidoptera of America north of Mexico. 

Trans. Am. Entomol. Soc. 11:245-337. 

1885. Miscellaneous notes on butterflies, habits of larvae, etc. oer Entomol. 

17:108-114. 

1887-1897. The butterflies of North America. Vol. II]. Am. Entomol. Soc, 

Philadelphia, Pennsylvania. [432] pp. 

1888a. Obituary of Annie M. Wittfeld. Canad. Entomol. 20:140. 

1888b. On Diadema misippus, Linn, in Florida. Canad. Entomol. 20:128. 

FRENCH, G. H. 1885. The butterflies of the eastern United States. J. B. Lippincott Co., 
Philadelphia, Pennsylvania. 402 pp. 

GROSSBECK, J. A. 1917. In Watson, F. E. (ed.), Insects of Florida IV. Lepidoptera. Bull. 
Am. Mus. Nat. His. 37: (Article 1):1—147. 

GROTE, A. R. 1882. Notes on certain Geometridae, with a new Byssodes from Florida. 
Papilio 2:100-101. 

1884. A new species of Nystalea. Papilio 4:7. 

HELLIER, W. R. 1965. Indian River, Florida’s treasure coast. Hurricane House, Pub- 
lishers, Inc., Coconut Grove, Florida. 128 pp. 

HOLLAND, W. J. 1898. The butterfly book. Doubleday, Page & Co., New York, New 
York. 382 pp. 

1903. The moth book. Doubleday, Page & Co., New York, New York. 479 pp. 

1931. The butterfly book, new and thoroughly revised edition. Doubleday and 
Co., Inc., Garden City, New York. 424 pp. 

HuLsT, G. D. 1886. Descriptions of new Pyralidae. Trans. Am. Entomol. Soc. 13:145- 
168. 


VOLUME 48, NUMBER 1 67 


KIMBALL, C. P. 1965. Arthropods of Florida and neighboring land areas. Vol. 1. Lep- 
idoptera of Florida. Div. of Plant Industry, Gainesville, Florida. 363 pp. 

KJERULFF, G. G. 1972. Tales of old Brevard. The South Brevard Hist. Soc., Melbourne, 
Florida. 121 pp. 

LANGLAIS, V. G. 1984. Cemetery census of Brevard County Florida, including Sebastian 
(Indian River County). Dist. by the author, Titusville, Florida. 155 pp. 

LINTNER, J. A. 1881. On some species of Nisoniades. Papilio 1:69-74. 

MAYNARD, C. J. 1891. A manual of North American butterflies. De Wolf, Fiske and 
Co., Boston, Massachusetts. 226 pp. 

MCALEENAN, J. 1991. Romantic horticulturist named M_[erritt] I.[sland] landmarks. 
Florida Today (19 Oct.):2A. 

MILLER, L. D. & F.M. BRown. 1981. A catalogue/checklist of the butterflies of America 
north of Mexico. Lepid. Soc. Memoir No. 2. 280 pp. 

Myers, R. L. & J. J. EWEL (eds.). 1990. Ecosystems of Florida. Univ. of Central Florida 
Press, Orlando, Florida. 765 pp. 

NEUMOEGEN, B. 1881. On a new species of Arctia from Florida. Papilio 1:9-10. 

SCUDDER, S. H. 1889. The butterflies of the eastern United States and Canada with 
special reference to New England. 3 Vols. Publ. by the author, Cambridge, Massa- 
chusetts. 1958 pp. 

SKINNER, H. 1894. Tachyris ilaire n. var. neumoegenii. Entomol. News 5:110. 

1896. Two new hesperids. Canad. Entomol. 28:187-188. 

1907. Rare butterflies. Entomol. News 18:22-24. 

1911. The larger boreal American Hesperidae, including Eudamus, Erycides, 

Pyrrhopyge and Megathymus. Trans. Am. Entomol. Soc. 37:169-209. 

1921. Atrytone kumskaka Scudder (Lep., Rhop.). Entomol. News 32:276-277. 

SKINNER, H. & R. C. WILLIAMS JR. 1924. On the male genitalia of the Hesperiidae of 
North America. Paper III. Trans. Am. Entomol. Soc. 49:129-153. 

1925. On the male genitalia of the Hesperiidae of North America. Papers [V- 
VI. Trans. Am. Entomol. Soc. 50:57-73, 141-156, 177-208. 

TEBEAU, C. W. 1980. A history of Florida. Univ. of Miami Press, Coral Gables, Florida. 
527 pp. 

TRUE, F. W. 1884. A muskrat with a round tail. Science 4:34. 

U.S. CENSUS BUREAU. 1900. Twelfth census of the United States, Brevard County, 
Florida, Precinct 13:184. 

WEIGLEY, R. F. (ed.). 1982. Philadelphia, a 300-year history. W. W. Norton & Co., 
New York, New York. 842 pp. 


Received for publication 1 May 1998; revised and accepted 9 August 1993. 


GENERAL NOTES 


Journal of the Lepidopterists’ Society 
48(1), 1994, 68-69 


MUSINEON TENUIFOLIUM (APIACEAE): NEW HOST OF FOUR 
NEBRASKA PAPILIO (PAPILIONIDAE) 


Additional key words: Artemisia dracunculus, Papilio machaon-complex, swallow- 
tail. 


The recorded larval host plants of four Papilio species, P. zelicaon nitra W. H. Edwards, 
P. indra Reakirt, P. polyxenes asterius Fabricius, and P. machaon bairdii (W. H. Ed- 
wards), encompass a variety of genera in three plant families, Asteraceae, Apiaceae, and 
Rutaceae. Here we report narrow-leaved musineon, Musineon tenuifolium Nutt. (Api- 
aceae), as a new larval host plant for these four swallowtails. 

Musineon tenuifolium is endemic to the west-central Great Plains, ranging from the 
Black Hills of South Dakota and Wyoming, south through eastern Wyoming and the 
Nebraska panhandle (Barkley 1977). Farrar (1990) characterizes M. tenuifolium as a 
native perennial favoring dry, rocky outcrops, growing from a caudex atop a thickened 
taproot. It possesses stemless, dissected leaves rising from a root crown to form a compact 
tuft of foliage. Small, cream to bright yellow flowers form compound umbels, approxi- 
mately 2.5 cm across, atop a leafless stalk normally less than 15 cm tall. In Nebraska, M. 
tenuifolium usually stays green the entire season except under severe drought conditions. 
It is an extremely local umbel, but often common in the appropriate habitat. 

M. tenuifolium was first brought to our attention in 1983 while collecting in the canyons 
of northwestern Sioux County, Nebraska. A search of M. tenuifolium foliage yielded 
larvae of both P. z. nitra and P. indra. M. tenuifolium is believed to be the exclusive 
host plant for these swallowtails in Nebraska, and supports a second brood of P. indra in 
our area. We also have observed larvae of P. z. nitra and P. indra feeding on M. 
tenuifolium in the Black Hills of South Dakota. 

On 20 June 1991, a collecting trip to Bull Canyon, Banner County, Nebraska resulted 
in about 40 swallowtail larvae collected on M. tenuifolium. Of these larvae, approximately 
one-half emerged as adults from mid-July through mid-August. Surprisingly, only three 
adults were P. z. nitra; others were P. p. asterius. 

While collecting in southern Scotts Bluff County, Nebraska on 25 May 1992, five 5th 
instar swallowtails were discovered on M. tenuifolium by the first author and Nick D. 
Theis. The larvae were removed and fed cut M. tenuifolium until pupation. Starting 15 
June 1992, pupae began to eclose. Emergent adults were P. m. bairdii, a species thought 
to have fed exclusively on silky wormwood, Artemisia dracunculus L. (Asteraceae), in 
Nebraska. 

These observations raise some intriguing questions regarding possible range extensions 
and the use of alternative host plants by members of the P. machaon-complex. It is likely 
that continued efforts of naturalists and collectors within and outside the state will lead 
to insights concerning these and subsequent questions. 

We thank Brett C. Ratcliffe and J. Ackland Jones for review of the manuscript. We 
are grateful to Neil E. Dankert and Jeff C. Germer, who assisted in collecting larvae. 
This is paper number 10264 of the journal series of the Nebraska Agricultural Research 
Division, University of Nebraska. This work was supported by the University of Nebraska 
Agricultural Experiment Station Project 17-055. 


LITERATURE CITED 


BARKLEY, T. E. 1977. Atlas of the flora of the Great Plains. Iowa State Univerin Press, 
Ames. 600 pp. 

FARRAR, JON. 1990. Field guide to wildflowers of Nebraska and the Great Plains. 
Nebraska Game and Parks Commission, Lincoln. 216 pp. 


VOLUME 48, NUMBER 1 69 


J. M. REISER, Department of Horticulture, University of Nebraska, Lincoln, Nebraska 
68583-0724, AND S. M. SPOMER, Department of Entomology, University of Nebraska, 
Lincoln, Nebraska 68583-0816. 


Received for publication 25 February 1998; revised and accepted 29 August 1998. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 69-71 


TWO NEW SYNONYMIES IN NEARCTIC EUCOSMA 
(TORTRICIDAE: OLETHREUTINAE) 


Additional key words: Eucosma wandana, E. uta, E. ustulatana, Kentucky, genitalia. 


In the course of identifying Eucosma specimens captured recently in Kentucky, we 
reached the conclusion that E. wandana Kearfott, E. uta Clarke, and E. ustulatana 
Blanchard & Knudson are one and the same morphospecies. Males captured at the same 
Kentucky locality on the same date match male holotypes of all three taxa. All specimens 
we examined that had been or could be assigned to these taxa have similar underlying 
forewing patterns varying in the proportion of dark brown to light orange brown. On 
average, males (n = 12) are darker brown than females (n = 44). 

Eucosma wandana was described from one male by W. D. Kearfott in 1907, but we 
also have seen a female with a “type” label not mentioned in the original description; E. 
uta was described from two males and one female by J. F. G. Clarke in 1953; and E. 
ustulatana was described from two males by A. Blanchard and E. C. Knudson in 1983. 
None of these short type series reveals the full range of color variation we observed in 
longer series. Moreover, Clarke seems to have overlooked E. wandana when he described 
E. uta; for differentiation, he used E. atomosana (Walsingham), a western species (Hein- 
rich 1928). In describing E. ustulatana, Blanchard and Knudson used E. wandana (as E. 
eumaea Meyrick) for comparison, but the diagnostic differences cited by them—smoother, 
more evenly convex anterior valval margin in E. ustulatana—can be ascribed to individual 
variation. No differences are discernible among holotype genitalia of E. wandana (as E. 
eumaea) (Heinrich 1923), E. uta (Clarke 1953), or E. ustulatana (Blanchard & Knudson 
1983). 

We summarize as follows: 


Eucosma wandana Kearfott 
(Figs. 1-3) 


Eucosma wandana Kearfott (1907) (holotype: male, Cincinnati, Ohio, 25 July [not August 
as in original description] 1904, A. F. Braun, in American Museum of Natural History 
[AMNH], New York, N.Y., wings illustrated in Fig. 1 here); Klots (1942). 

E. eumaea E. Meyrick (1912); Heinrich (1923) (holotype genitalia photo-illustrated); Klots 
(1942) (invalid replacement name). 

E. uta Clarke (1953) (holotype: male, Putnam Co., Illinois, 10 July 1939, M. O. Glenn, 
in National Museum of Natural History [NMNH], Washington, D.C.; male and female 
genitalia sketch-illustrated, wings of paratype photo-illustrated); Godfrey, et al. (1987) 
wing of female photo-illustrated. New Synonymy. 

E. ustulatana Blanchard & Knudson (1983) (holotype: male, Brenham, Washington Co., 
Texas, 4 June 1979, E. C. Knudson, genitalia prep. WEM 36924, in NMNH, wings 
photo-illustrated, male genitalia of paratype photo-illustrated). New Synonymy. 


Eucosma wandana is probably multivoltine. Capture dates range from 4 June to 15 
September (n = 53). The known distribution is Kentucky, Ohio, Illinois, Missouri, Arkansas, 
Kansas, Georgia, Florida, and Texas. Forewing length of males ranges from 7.5 to 8.5 
mm (n = 12); of females, 7.0 to 9.25 mm (n = 44). Eucosma wandana is one of at least 
11 eastern species of Eucosma having ventral extensions of female papillae anales (Fig. 


70 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


3 


Fics. 1-3. Eucosma wandana. 1, Wings of holotype. 2, Genitalia of male. 3, Genitalia 
of female. Z 


3), an accessory whose function is not yet known (Miller 1987). The larval food plant is 
unknown. 

We thank F. H. Rindge, G. L. Godfrey, K. R. Methven, J. R. Heitzman, and D. Profant 
for specimen loans. 

Material examined. We examined specimens as follows, labeling all except primary 
types “Voucher, Gibson & Miller 1992”: ARKANSAS: Devil’s Den St. Pk., Washington 
Co., one male, 6.VII.66; one female, 6.VII.66 (genit, prep. USNM 70317); one female, 
26. VI.66; one female, 3. VII.66 (genit. prep. USNM 70316) all R. W. Hodges; Washington 
Co., “type”, female. KANSAS: Pittsburg, E. L. Todd, one female, 28.VI.54; one female, 
2.VII.54. GEORGIA: Atlanta, P. W. Fattig, one male, 31.VII.41. TEXAS: E. ustulatana 
holotype; Dallas, “714”, Fernald collection, one female (all foregoing in NMNH). OHIO: 
E. wandana holotype (in AMNH). KENTUCKY: Campbell Co., Jolly Co. Pk., three 
females, 15. VII.82 (female genit. prep. LDG 134) (Fig. 3); Gallatin Co., Markland Dam, 
two females, two males, 3. VIII.88 (male genit. prep. LDG 048) (Fig. 2); Boone Co., Big 
Bone Lick St. Pk., one female, 7.VII.81; one female, 27. VII.89; Owsley Co., near Boone- 


VOLUME 48, NUMBER 1 7a 


ville, two females, two males, 22.VI.84 (male genit. prep. LDG 047); Bullitt Co., near 
Shepherdsville, one female, 8.VII.88 (all L. D. Gibson, in L. D. Gibson collection). MIS- 
SOURI: Cape Girardeau, one female, 25. VII.78; Independence, one male, 30. VI.76; Clay 
Co., Coolie Lk., one female, 18.VII.68; one female, 22. VII.72 (all J. R. Heitzman, in J. 
R. Heitzman collection). ILLINOIS: E. uta holotype; Oconee, two females, 8-15. VII; one 
female 1—7.VII (female genit. prep. USNM 70670); Decatur, one female, 8-15.VII; Put- 
nam Co., M. O. Glenn, one female, 30. VII.74; one male, 18. VII.73; one female, 3. VIII.65; 
one male, 25.VII.74 (male genit. prep. WEM 176923); one female, 13.VIII.43 (female 
genit. prep. USNM 70320) (E. uta paratype); (all in NMNH); Putnam Co., M. O. Glenn, 
one male, 23.VI.56, “5057”; one male, date missing, “5157”; one female, 5.VII.53; one 
female, 24. VII.68; one female, 11.VIII.74; one female, 15.VII.64; one female, 5. VIII.61 
(abdomen missing); one female, 28.VII.64; one female, 2.VII.56; one female, 15.1X.54; 
one female, 28. VII.56; one female, 21. VII.56; one female, 23. VII.73; one female, 10. VII.39; 
one female, [illegible]. VII.65; one female, 17.VII.64; one female, 29.V1I.64; one female, 
5. VIII.47 (E. uta paratype) (all in Illinois Natural History Survey Collection). FLORIDA: 
Liberty Co., Torreya St. Pk., one female, 7.VII.88 (female genit. prep. LDG 131) (D. 
Profant, in D. Profant collection). 


LITERATURE CITED 


BLANCHARD, A. & E. C. KNUDSON. 1983. New North American species of Eucosmini 
(Lepidoptera: Tortricidae). Proc. Entomol. Soc. Wash. 85:845-852. 

CLARKE, J. F. G. 1953. New species of Olethreutidae from Illinois (Lepidoptera). J. 
Wash. Acad. Sci. 43:226-231. 

GopFREY, G. L., E. D. CaAsHaTT & M. O. GLENN. 1987. Microlepidoptera from the 
Sandy Creek and Illinois River region: An annotated checklist of the suborders 
Dacnonypha, Monotrysia, and Ditrysia (in part) (Insecta). Il]. Nat. Hist. Surv. Spec. 
Pub. 7. 44 pp. 

HEINRICH, C. 1923. Revision of the North American moths of the subfamily Eucosminae 
of the family Olethreutidae. U.S. Natl. Mus. Bull. 123. 298 pp. 

KEARFOTT, W. D. 1907. New North American Tortricidae. Trans. Am. Entomol. Soc. 
33:1-98. 

Kiots, A. B. 1942. Type material of North American microlepidoptera other than 
Aegeriidae in the American Museum of Natural History. Bull. Am. Mus. Nat. Hist. 
79:391-424. 

MEYRICK, E. 1912. On some impossible specific names in micro-Lepidoptera. Entomol. 
Mon. Mag. 23:32-36. 

MILLER, W. E. 1987. Guide to the olethreutine moths of Midland North American 
(Tortricidae). U.S. Dept. Agr. Handb. 660. 104 pp. 


LORAN D. GIBSON, 8496 Pheasant Drive, Florence, Kentucky 41042, AND WILLIAM 
E. MILLER, Department of Entomology, University of Minnesota, St. Paul, Minnesota 
55108. 

RECEIVED FOR PUBLICATION 28 JANUARY 1993; REVISED AND ACCEPTED 27 JULY 1993. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 71-78 


ADDITIONAL DATA ON THE GEOGRAPHICAL DISTRIBUTION AND 
ADULT ACTIVITY OF THE DIURNAL, MIMETIC PLUME MOTH, 
OIDAEMATOPHORUS CHAMELAI (PTEROPHORIDAE) 


Additional key words: Costa Rica, Mexico, Croton, Cordia. 


An extraordinary, black pterophorid with enlarged scale tufts on the hind legs was 
described from Mexico by Gielis (1992), but neither its strikingly wasp-like appearance 


2 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


nor its diurnal behavior, unusual for the family, were emphasized. In addition to the type 
series of seven specimens, most of which were collected by Chemsak in 1985-86, we have 
taken a larger number in more recent years, documenting a more extensive flight period. 
Moreover, two specimens from Costa Rica were erroneously recorded from Mexico by 
Gielis. 

Geographical distribution. Oidaematophorus chamelai has been recorded along the 
immediate Pacific coast of Middle America, in Mexico near Elota in central Sinaloa, and 
Chamela, Jalisco, and in northwestern Costa Rica in Guanacaste Province. An inland 
occurrence in Mexico, indicated on the mapped distribution by Gielis in the vicinity of 
Guadalajara, evidently was based on his misinterpretation of the label on specimens from 
Costa Rica (“C.R.: Playa Tamarindo, Guan. Prov.’’) as the State of Guanajuato, Mexico. 
A specimen from south of Elota, Sinaloa, was illustrated and cited but not mapped. 

Seasonal occurrence. In Mexico we have taken adults of O. chamelai during each 
month that we have visited the Estacion de Biologia Chamela: July, August, September, 
October, December; while the Costa Rica specimens were collected in June. Whether or 
not the species flies during the dry season (December—June in Mexico, January—April in 
Costa Rica) cannot be determined from our sampling. 

Mimetic appearance and diurnal activity. Oidaematophorus chamelai and two closely 
related South American species, O. ochracealis (Walker) and O. paraochracealis Gielis, 
differ markedly from typical members of the genus in having the wings relatively broad 
and uniformly black with purplish iridescence or “brown-black”’ (Gielis 1992). Oidae- 
matophorus chamelai is particularly wasp-like, an image enhanced by broad, cream- 
colored, dorsolateral spots on abdominal segments 2 + 3 that render the appearance of 
the constricted petiole of many ichneumonids and sphecids, and by seemingly aposematic 
red tegulae and large yellowish “eyespots” of the metathorax. Another pair of cream 
colored dorsal spots at the base of segment 8, which were lacking on the Sinaloa female 
Gielis illustrated, are variable from a trace to conspicuous in both sexes. The male has 
long, white, eversible hair brushes flanking the genitalia that were not mentioned by 
Gielis. The enlarged scale tufts of the hind tibiae (misinterpreted as the first tarsal segment 
in the original description) and tarsi in the male and tarsi only in the female are reminiscent 
of many ctenuchine Arctiidae, such as Macrocneme, that are believed to be members of 
mimetic assemblages. D. L. Janzen has reared one species of Macrocneme and two of 
the closely related Poliopastea from Mesechites trifida (Apocynaceae), a plant family 
well known for its cardiac glycocides. The diurnal, aposematic larvae and adults of these 
two genera are superficially quite similar and presumably are distasteful to vertebrate 
predators (Janzen in litt., Dietz 1994). 

While nearly all Nearctic and Neotropical pterophorids that we have encountered are 
nocturnal, O. chamelai (and presumably the other species of the O. ochracealis complex) 
are primarily diurnal. We have observed adults of O. chamelai at flowers of three shrub 
and tree species of Croton (Euphorbiaceae) at the Estacion de Biologia Chamela. Their 
time of activity corresponded with that of other flower visitors such as cerambycid and 
scarabaeid beetles, wasps, bees, and hesperiids. In Costa Rica, males were collected by 
sweeping inflorescences of an arboreal Cordia (Boraginaceae), 5-6 m above the ground, 
employing an extensible tropical net. In addition, however, about 7 individuals of 17 
taken at Chamela were attracted nocturnally to ultraviolet or white lights. 

Location of specimens. The holotype is housed in the Essig Museum of Entomology, 
University of California, Berkeley (UCB), along with later collections. Specimens also are 
deposited in the Instituto de Biodiversidad, Santa Domino de Heredia, Costa Rica (INBio); 
Instituto de Biologia, Universidad Nacional Autonoma de México, Mexico City (UNAM); 
Los Angeles County Museum of Natural History (LACM); U. California, Riverside (UCR); 
and the U. S. National Museum of Natural History, Washington, D.C. (NMNH). The 
type series was erroneously cited as LACM; the specimens had been on loan to J. P. 
Donahue from UCB and UCR and were so labelled when forwarded to Gielis, but their 
ownership was not communicated. 

Material examined. COSTA RICA: Playa Tamarindo, Guan. Prov., 2 6, VI-14-86, 
“arboreal Croton fls. 15-18 ft. above ground, tropical net’’ (Chemsak, H: Katsura, A. & 
M. Michelbacher). MEXICO: Estacion de Biologia Chamela, Jalisco, ¢ Holotype, ? VII- 


VOLUME 48, NUMBER 1 73 


8/16-85 (Chemsak, Katsura, Michelbacher); 2 4, 2 X-13/23-86 (Chemsak); 2 VII-12/15- 
87 (Chemsak & E. G. Linsley); 2 2 X-16/19-87, 6 X-21/22-87, blacklight (Chemsak & 
Powell); 6 [X-27/28-88, blacklight (P. A. Opler); 2 XII-1-88 (Chemsak); 2 6, 2 VII-23-90, 
1 6 at light (Chemsak); ° VII-13-92, 6 VII-20-92, at lights (Chemsak); 1 6, VIJ-15-93, at 
Croton flowers (B. Eya). 

We thank D. H. Janzen for unpublished observations on ctenuchid biology in Costa 
Rica and J. P. Donahue for review and comments on the manuscript. 


LITERATURE CITED 


Dietz, R. E. Ill. 1994. Systematics and biology of the genus Macroeneme Hiibner 
(Lepidoptera: Ctenuchidae). Univ. Calif. Publ. Entomol. In press. 

GIELIS, C. 1992. Neotropical Pterophoridae 6. The Oidaematophorus ochracealis com- 
plex (Lepidoptera). Phegea 20:81—94. 


J. A. POWELL AND J. A. CHEMSAK, Essig Museum of Entomology, University of 
California, Berkeley, California 94720. 


Received for publication 6 April 1993; revised and accepted 4 September 1993. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 74-76 


BOOK REVIEWS 


KEYS TO THE INSECTS OF THE EUROPEAN PART OF THE USSR (G. S. Medvedev, chief 
editor). VOLUME IV (LEPIDOPTERA), PART II, by M. I. Falkovitsh (ed.) et al. 1990. E. J. 
Brill, Leiden. (translation of: OPREDELITEL NASEKOMYKH EVROPEISKOI CHASTI SSSR, 
TOM IV, CHESHUEKRYLYE, VTORAIA CHAST. Nauka Publishers, Leningrad, 1981—+trans- 
lator: B. R. Sharma). x + 1092 pp., 675 figs. Hard cover, 16 x 24 cm, ISBN 90-04-08926- 
8. $160.00 U.S. Available from E. J. Brill (U.S.A.) Inc., 24 Hudson Street, Kinderhook, 
New York 12106. 


This remarkable, bulky handbook is the second part of a work devoted to the Lepi- 
doptera of western Russia. The English version of Part One was issued in 1987 (Amerind 
Publishing Co. Pvt. Ltd, New Delhi—edition supervised by the U.S.D.A., Washington, 
D.C.), and it covered the non-ditrysian families (with only a superficial treatment of the 
Nepticulidae), and eight families among the lower Ditrysia, namely the Psychidae and 
all members of the Zygaenoidea, Cossoidea, Sesioidea and Tortricoidea. Part Two deals 
with a larger number of families (29 if one accepts the classification that I proposed in 
1991: see Entomol. Scand. 22:90-91). These are the Eriocottidae and Tineidae (including 
the “Euplocamidae”’ and “Hieroxestidae’’), all members of the Gracillarioidea, Ypono- 
meutoidea, Choreutoidea, Urodoidea (Wockia Heinemann: p. 508, as a “plutellid” genus), 
Schreckensteinioidea, and Epermenioidea, and all the gelechioid families with the ex- 
ception of most Coleophoridae (only the Amphisbatinae being treated: pp. 792 and 801, 
among the Oecophoridae sensu auct.). Parts One and Two thus provide keys to the 
“Micromoths” of nearly all the families known to occur in the European part of Russia. 

The present book is firmly bound in boards, nicely presented, and has a good quality 
paper, i.e., opaque and whiter than that of the Russian edition. Compared with the latter, 
the book is thicker (approximately 7 cm) and of a different format: 15.5 by 24 cm instead 
of 17.5 by 27 cm. Most figures, however, have not been reduced and are as good as those 
of the original publication. Only a few figures are less satisfactorily printed, such as 
Figures 191 and 193. The page numbers of the Russian original are mentioned in the 
lefthand margin, and thus often precede the numbers of the figures (which may be a bit 
confusing). The arrangement of the figures is usually rather practical, although it should 
have been slightly different in a few cases: for example, Figures 11 and 12 (Euplocamus 
Latreille) are found on page 16, in the “Key to suborders and families,” whereas they 
correspond to the “Euplocamidae”’ text (pp. 24-25); Figures 499 to 505 (Schreckensteinia 
Hubner and Heliodines Stainton) should have been placed between the Schreckenstei- 
niidae (pp. 697-698) and Heliodinidae (pp. 699-700), rather than at the end of the 
Stathmopodidae chapter; etc. 

On the whole, the English translation appears quite faithful, and no changes were 
introduced, either nomenclatural or taxonomic. That is definitely a reasonable option, 
although typographical errors might have been rectified for certain Latin names. For 
instance, Bucculatrix gnaphaliellea (p. 193), B. frangulella (p. 195), Caloptilia populo- 
torum (p. 252), Milliereia (p. 414), Mompha propinguella (p. 704), Pseudatemella (p. 
792), and Oliaria (p. 1001), are all incorrect spellings for, respectively, Bucculatrix gna- 
phaliella (Treitschke), B. frangutella (Goeze), Caloptilia populetorum (Zeller), Millieria 
Ragonot, Mompha propinquella (Stainton), Pseudatemelia Rebel, and Uliaria Dumont. 

Nine authors contributed to Part Two, namely: A. S. Danilevsky (Choreutidae), M. I. 
Falkovitsh (Heliodinidae, Scythridae, Stathmopodidae, Schreckensteiniidae, Epermeni- 
idae), Z. S. Gershenzon (Yponomeutidae), V. I. Kuznetzov (Gracillariidae, Glyphipteri- 
gidae, Choreutidae), A. L. Lvovsky (Oecophoridae sensu auct.), V. I. Piskunov (Sym- 
mocidae, Lecithoceridae, Blastobasidae, Gelechiidae), S. V. Seksajeva (Bucculatrigidae, 
Gracillariidae, Phyllocnistinae, Lyonetiidae), S. Yu. Sinev (Elachistidae, Batrachedridae, 
Momphidae, Cosmopterigidae), and A. K. Zagulajev (Tineoidea, Douglasiidae, Roesler- 
stammiidae, Urodidae, and several yponomeutoid and gelechioid families). Most of these 
entomologists are well known specialists of the families in question, so that the proposed 
keys are usually quite reliable. Sometimes they could not examine a number of foreign 
(European) species, their work in such cases being largely based on that of other specialists 


VOLUME 48, NUMBER 1 To 


(e.g., R. Gaedike’s papers on Douglasiidae and Epermeniidae). Much in the same way, 
the drawings were made by several artists or even, sometimes, by certain of the above- 
mentioned lepidopterists. These illustrations range from fair to excellent, and are often 
original, though explicitly taken from other works in several cases. I would like to em- 
phasize the abundance of the figures provided throughout the book: nearly 530 drawings 
represent adults or their wing pattern; nearly 240, wing venation schemata; about 1700 
and nearly 470, male and female genital structures, respectively; and so on. As in Part 
One, photography was not used, probably because diagnostic features may be more easily 
distinguishable with line drawings. 

After a short abstract (p. v), a preface (pp. vii-viii) and a table of contents (pp. ix—x), 
the handbook starts with a key to suborders and families (pp. 1-23). Elaborated by M. I. 
Falkovitsh and A. K. Zagulajev, this key is almost identical with that published in Part 
One, except for minor changes: the translation is better, abbreviations have been avoided, 
and illustrations are slightly more numerous. Various parts of the key are interesting and 
original, but a few remarks have to be made: 


* many of the characters selected for a given family cannot be generalized on a 


worldwide scale 

according to the seventh alternative, tibial spurs would be absent from the Hepialidae, 
but they are actually present in genus Gazoryctra Hiibner, one species of which is 
treated in Part One (as a member of Korscheltellus Borner) 

* alternatives 13 and 22 are chiefly based on a slight difference in the “breadth” of 
the head, not on a clear-cut demarcation between two traits 

several inaccuracies must be corrected: ocelli are present in many Eriocottidae (con- 
trary to alternative 16 statement), the maxillary palpi may be well developed in 
Adelidae since Nematopogon Zeller definitely belongs to this family (90), the Sesiidae 
possess a long frenulum (118), tympana are distinctly present in the Thaumetopoeinae 
(Notodontidae) (182), ete. 

* “Endromidae’” has been omitted after “In forewings R with 4 branches (R2-R5) 
stalked together” (alternative 190); in the description of the Nolidae (alternative 
202), “ocelli” obviously represents a slip, and should be replaced by “compound 
eyes 

the nomenclature is wrong or outmoded for certain groups: in particular, Brachodidae 
must be substituted for “Atychiidae,’’ Saturniidae for “Attacidae,” and Thyatirinae 
(Drepanidae) for “Tetheidae” 

several “families” have to be downgraded to subfamily rank (or even to tribe rank): 
“Phyllocnistidae,” “Ethmiidae,”’ ““Galleriidae,” “Phycitidae,” “Pyraustidae,”’ “Lith- 
osiidae,’ “Ctenuchidae,” and so on. 


* 


The following section deals with about 30 families, for which are provided keys for 
the identification of genera and species (pp. 24-1024). For each family, one can find a 
general diagnosis and at least one illustrated key to species, based on external characters 
and genitalia. Frequently, there are two separate keys to species (one being based on 
external characters; the second, on male genitalia), and there may also be a third one, 
based on female genitalia. In addition, there is a key to genera (or to subfamilies and 
genera), unless the family contains only one genus in western Russia. By and large, all 
these keys are more accurate than the one to suborders and families. Although the book 
is not intended as a taxonomic revision, it provides useful data for each species (e.g., 
distribution and larval food plants), and includes some interesting proposals or suggestions: 
Eretmocera Zeller, often misplaced in recent literature, is rightly placed within the 
Scythridae (“Scythrididae’’); in the Oecophoridae sensu auct., Anchinia Hibner follows 
immediately Hypercallia Stephens, probably because A. L. Lvovsky regards these genera 
as close relatives, a point of view in agreement with my recent reappraisal of the gelechioid 
classification (I have transferred both genera to the Elachistidae Hypertrophinae: Alexanor 
16:239-255); Uliaria Dumont is rightly considered a member of the Gelechiidae Di- 
chomerinae (the genus has been occasionally allocated to the “Autostichinae’’); etc. Of 
course, the adopted classification and nomenclature would also require a number of 
amendments: Odites Walsingham is definitely not a member of the Xyloryctidae [see, 


76 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


e.g., Hodges 1978: Moths Am. N. of Mexico 6(1):8-9]; Millieria Ragonot belongs to the 
Choreutidae, not to the Glyphipterigidae (see, e.g., Heppner 1982: Smithson. Contr. Zool. 
370:1-27); Phyllonorycter Hiibner must be substituted for Lithocolletis Hiibner, Ypso- 
lopha Latreille for Ypsolophus Fabricius, Lecithocera nigrana (Duponchel) for Lecithoce- 
ra luticornella (Zeller), and so on. 

A list of references (pp. 1025-1041) and an index of Latin names (pp. 1061-1092) 
conclude the book, along with a list of abbreviations of authors (pp. 1042-1043), and a 
useful list of botanical names, which gives the correspondence between common and 
Latin denominations (pp. 1044-1060). The present handbook is highly valuable because 
it includes a large portion of the European fauna of “Micromoths,” for which no equiv- 
alent, recent work has been published up to now. For instance, it takes into account a 
high percentage of the species known to occur in France: nearly 80% for Bucculatrix 
Zeller, more than 90% for Caloptilia Hubner and Ypsolopha Latreille, 90% for Ethmia 
Hubner, about 60% for the Scythridae, 93% for the Momphidae, 100% for the Choreutidae, 
etc. 

In conclusion, this work should be included in the private library of any microlepi- 
dopterist interested in the Palaearctic fauna. It is to be wished that a similar translation 
will soon be available for Part Three (dealing chiefly with Pyrales and Plume Moths). 


JoEL MINET, Muséum national d Histoire naturelle, Entomologie, 45 rue Buffon, 
F-75005, Paris, France. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 76-77 


BUTTERFLY GARDENING IN THE SOUTH: CULTIVATING PLANTS THAT ATTRACT BUTTER- 
FLIES, by Geyata Ajilvsgi. 1990. Taylor Publishing Company, Dallas, Texas. xi + 348 pp., 
196 color photographs, 2 line drawings and 7 diagrams. Hard cover, 20.3 em x 28 cm, 
ISBN 0-87833-738-5. $34.95. 


Novice and master gardeners in the southern U.S., particularly in the Rio Grande 
Valley, Texas, have an extraordinary treat in store with this volume. Ms. Ajilvsgi inves- 
tigates every aspect of a butterfly garden, from her brief introduction on the significance 
of butterflies and their means of survival to appropriate commercial and private sources 
for obtaining seed and other garden supplies. The brief forward by Chess Ezzell McKinney, 
Chairman, Preservation of Butterflies, National State Garden Clubs, sets the stage for this 
treasury of information, and, although the title focuses on butterfly gardening, the em- 
phasis is on gardening with a capital “G.” 

There are special sections of the book devoted to creating a personal butterfly garden 
and to methods of attracting butterflies, highlighted by personal observations on such 
topics as the important characteristics of floral nectaries (color, shape and fragrance) and 
how to choose the appropriate plants, with one of my favorites—adopt a weed. Two 
other chapters detail the actual planning and planting of a butterfly garden. For the 
novice, or for butterfly watchers with limited space, there is the “instant” butterfly garden. 
For example, a fence, porch, or wall can accommodate hanging baskets or vines, and 
even the edges of driveways or window boxes can function as butterfly gardens. Special 
attention is given to the selection of plants, maintenance of the garden, and even to the 
introduction of caterpillars and chrysalids. For those lepidopterists with a grander vision 
in mind, a series of diagrams (pp. 78-83) provides garden plans for almost any geographic 
setting in the south. This thorough chapter on planning includes discussions of soils, 
preparation of the flower beds, and selection of appropriate plants, including native 
species, in addition to some forethought about flight patterns through the garden and 
appropriate areas for water and puddling stations. A chapter on “butterfly-friendly pest 
controls” emphasizes biological and physical controls, companion and repellent plants, 
and natural insecticides. Butterflies of south Texas, and especially of the Rio Grande 


VOLUME 48, NUMBER 1 Tt 


Valley, are featured in the special south Texas garden, which lists local butterflies and 
their associated larval hostplants based on the current literature and on the author’s own 
field observations. 

Detailed analyses for 50 butterfly species that are easily visible in the field or backyard 
are included and organized according to the common name followed by the scientific 
name. Each diagnosis generally includes a color illustration, size, range, flight period, 
number of broods, overwintering stage, and brief descriptions of the adult, egg, and larva, 
and of the hostplant and parts of the plant consumed. Curiously, descriptions of the 
chrysalis are omitted. Favorite nectar sources, their bloom period, height class, cultivation, 
and other notes are also listed. Descriptions of adult behavior, especially favorite perches 
and interaction with nectar sources, are interspersed throughout this section. 

Separate chapters treat larval hostplants and nectar resources, with the plants subdivided 
according to type (trees, vines, and herbs). These chapters feature both cultivated and 
native species with a special perspective on south Texas butterflies and with descriptions 
and accompanying color illustrations for 40 plants in each category. The chapter on larval 
hostplants provides information on their cultivation, associated larval taxa, and other 
notes. A longer list of all potential host plants gives the common name of their associated 
butterflies, the normal plant range, and the geographic area in which these plants are 
located in Texas (pp. 209-242). Similarly, the chapter on nectar sources lists the most 
appropriate yet common plants, their habitat association, and identification as a larval 
hostplant, if appropriate. An addendum list of nectar plants (pp. 281-311), including 
both cultivated and native naturalized species, is subdivided annuals versus perennials 
(herb), with special annotation indicating height, color and bloom period. 

A wealth of other information can be found in this book, including basic advice and 
tips on the photography of butterflies. Other features include maps of U.S. hardiness zones 
and of the regional subdivisions of Texas considered. A compendium of appendices lists 
the major butterfly and plant organizations and societies and their associated publications, 
butterfly gardens and special events, and garden magazines and newsletters with articles 
on butterfly gardening. Other lists provide sources for additional information on habitat 
preservation, garden seeds, and supplies. The diverse bibliography supplies references on 
butterflies and other insects; garden, plant and wildflower guides; and photography. The 
butterfly and plant index is consistent in format, listing species by common name followed 
by the associated scientific name. 

This book is exceedingly well researched and delightfully written. It is refreshing to 
read the author's personal observations on gardening and butterflies, especially those on 
behavior. For the taxonomist, the organization of the butterfly diagnoses is a little dis- 
concerting, with, for example, the Janais Patch (Chlosyne janais) and the Theona Check- 
erspot (Thessalia theona) interspersed among representatives of the Hesperiidae. Similar 
problems occur in the Lycaenidae. 

Although the use of common names as the main reference point may be attractive to 
introduce the uninitiated gardener to butterflies, such names vary in use from one section 
of the country to another, leading to confusion. Thus, the Gray Calico and Gray-skirted 
Calico refer to the same beast, Hamadryas februa. One further minor hurdle concerns 
the type size of the index (6 pt.), which is generally in vogue for curatorial staffs but may 
be a little small for the average reader. 

These few problems notwithstanding, this book is beautifully illustrated and provides 
a treasury of vital information on butterfly gardening. For anyone even remotely interested 
in southern gardening and butterflies, this book is absolutely essential reading. 


JACQUELINE Y. MILLER, Allyn Museum of Entomology, Florida Museum of Natural 
History, 3621 Bay Shore Road, Sarasota, Florida 34284. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 78-79 


BUTTERFLY GARDENING: CREATING SUMMER MAGIC IN YOUR GARDEN, created by the 
Xerces Society in association with the Smithsonian Institution. 1990. Sierra Club Books, 
San Francisco, and National Wildlife Federation, Washington, D.C. xv + 192 pp., 118 
color photographs, 4 diagrams. Softcover, 18.5 x 23.5 cm, ISBN 0-87156-615-X. $18.95. 


The cooperative spirit of this volume is reflected not only in the educational and 
conservation organizations involved in its production, but also by the wealth of individ- 
uals—authors, naturalists, and photographers—who so generously donated their time and 
efforts. Butterfly Gardening is dedicated to the broad scale conservation of butterflies 
and their native food plants. It also celebrates the diversity of nature and encourages all 
lepidopterists to observe ecology in action. With the loss of native habitats, there also has 
been a noticeable disappearance of butterfly species, which serve as indicators of major 
changes in ecosystems. 

A brief Introduction by Dennis Murphy focuses on some of the problems in butterfly 
conservation, suggests possible solutions, and sketches the contributions of the other nine 
authors of the book. Such a conservation approach to butterfly gardening would be 
incomplete without a contribution from the Honorable Miriam Rothschild, the eminent 
entomologist who has devoted so much time to the study of butterflies and other inver- 
tebrates. Dr. Rothschild’s keen insight has made naturalists more aware of conservation 
and of the significant role of wildflowers in ecosystems. Dr. Rothschild wrote three chapters 
for this book. The first describes the joys and challenges of Gardening with Butterflies; 
the second explores The Visual Perception of Lepidoptera and the importance of pro- 
viding appropriate nectar sources in butterfly gardens; the third describes The Life Cycle 
of the Large White Butterfly. Some life history aspects crucial to the survival of butterflies 
and moths in nature—metamorphosis, mate recognition, migration, temperature regu- 
lation, natural predators, and defense through camouflage and cryptic coloration and 
mutualism—are discussed by Dave Winter in his erudite chapter (The Struggle to Sur- 
vive). From a more personal perspective, Jo Brewer recounts her experiences as a butterfly 
gardener—providing descriptions of some of her more spectacular triumphs, sprinkled 
with the inevitable problems she encountered (Notes from a Butterfly Gardener). She 
also evaluates the propriety of introducing butterfly species into new habitats and discusses 
the use of native plant species versus exotic species in butterfly gardens. 

Mary Booth, a landscape architect, with supporting text by Melody Mackey Allen, 
provides butterfly garden designs with emphasis on color. Four basic arrangements from 
simple to more sophisticated plans are provided and suggestions for flowering plants are 
included. Some do’s and don'ts with regard to conservation and to introduction of plant 
species that may be difficult to control are also considered. This chapter (Butterfly Garden 
Design) concludes with a master plant list of 30 flowering plants that are common nectar 
sources and, in some cases, larval hostplants. This annotated list provides a description of 
the type (shrub versus hardy perennial), height, color, bloom season, exposure, and soil 
requirements for each species. The food requirements for moths are not neglected owing 
to the thoughtful inclusion of Dave Winter’s chapter on nocturnal nectar sources (Moths 
and the Garden at Night). 

Conservation issues concernign habitat are addressed in the chapter Wildflowers in 
the Planned Landscape by David Northington. As the Executive Director of the National 
Wildflower Research Center, he discusses the disappearance of plants and the resulting 
ecological consequences for animal species. The poignant discussion by Stanwyn Shetler 
in Butterfly Gardening and Conservation addresses these subjects from a naturalist’s 
viewpoint and delves further into the importance of plant/insect interactions. Dr. Shetler 
actively supports increased public education and awareness of conservation efforts by 
promoting gardening to teach preservation, rehabilitation, and restoration of diverse 
natural habitats. 

The final chapters offer suggestions for enhancing personal enjoyment of butterfly 


VOLUME 48, NUMBER 1 79 


gardens, with excellent tips on close-up photography and butterfly observations by the 
incomparable Edward Ross (Enriching Your Personal Landscape and Butterfly Photog- 
raphy) and additional intriguing ideas for butterfly watching by Robert Michael Pyle. 
Following a short Afterword by Pyle are two appendices: a list of nectar plants for North 
American butterflies and moths and an annotated list of the most familiar North American 
butterflies and their larval food plants. Resource lists for obtaining garden plants and 
entomological equipment, a lengthy bibliography, and an index to both scientific and 
common names complete this useful volume. 

An attractive glossy cover adds an exquisite touch to this extraordinarily illustrated 
volume. Superbly written, this book offers an excellent mixture of conservation philosophy 
and biological common sense with a focus on butterflies and moths. With such a com- 
pilation of authors, the book is anecdotal, and if there is any fault, it is that parts of some 
chapters may appear redundant. However, such duplicate treatment is refreshing, inas- 
much as different perspectives reflect the diversity of thinking on various subjects, such 
as appropriate nectar sources and problems encountered in rearing species. The Xerces 
Society and Smithsonian Institution are indeed to be commended for producing this 
volume. Through conservation, restoration, and management of native habitats initiated 
at the backyard level, we can increase public awareness of how complex yet fragile nature 
is and make a new beginning at protecting Lepidoptera. 


JACQUELINE Y. MILLER, Allyn Museum of Entomology, Florida Museum of Natural 
History, 3621 Bay Shore Road, Sarasota, Florida 34234. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 79-80 


THE ONTARIO BUTTERFLY ATLAS, by Anthony M. Holmes, Quimby F. Hess, Ronald R. 
Tasker, and Alan J. Hanks. 1992. Toronto Entomologists’ Association, 34 Seaton Drive, 
Aurora, Ontario L4G 2K1, Canada. 167 pp., 13 color plates. Soft cover, 20.5 x 25.5 cm, 
ISBN 0-921631-11-1. Available from the Association for $29 Canadian (including taxes 
& p/h) or $26 U.S. (including p/h) (no Master-Card or Visa). 


This attractively covered paper-back culminates the efforts of the Toronto Entomol- 
ogists Association (TEA) by recording the skippers and butterflies found in Ontario. The 
title may be somewhat misleading, however, as this publication is more of an annotated 
checklist of Ontario Rhophalocera. 

The preface, brief author backgrounds, and table of contents are followed by an 


introduction that describes the purposes of the Atlas: “to summarize ... the distribution 
and some characteristics of Ontario butterflies .. . , to encourage . . . others to explore the 
distribution ..., and as a reference for planning efforts to conserve ... rare species. ...” 


This introduction also includes information on TEA, nomenclature, timetables, habitat, 
status (employing symbols of occurrence used by the Nature Conservancy of Canada), 
records, figures showing counties and districts, physiographic features (Hudson Bay low- 
lands to deciduous forest region in southwestern Ontario), life zones (Hudsonian, Cana- 
dian, Transition and Carolinian), and problem species. The latter category includes several 
butterflies treated as species by some authors and as subspecies by others; also, because 
some species of Erynnis and Satyrium are difficult to identify, some records may be 
inaccurate. 

The main portion of the Atlas treats 138 species, including Basilarchia arthemis ar- 
themis and B. a. astyanax as separate entities. The authors make no attempt to differentiate 
subspecies except in a few cases. Each family is introduced with beautiful photographs 
of adults in natural settings or poses, with the exception of the Hackberry Butterfly on 
page 111 that suspiciously resembles a pinned specimen! 

The authors devote a full page to each species, with brief notes on timetable, including 
broods and “hibernates,”’ and occurrence, including habitat, food plant, distribution, and 


80 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


status. Also included on the species page is an occurrence calendar-graph for each stage 
of the species and a map of Ontario showing distribution by dots placed in each county 
and district of occurrence. I wish the authors had included more biological information 
and “characteristics” of each species instead of devoting more than half a page to each 
distribution map. The 12 color plates include 2 habitat photographs and 27 individual 
photographs that show 7 larvae and pupa and 20 adults, representing 22 species. 

Following the main species section is information on 19 stray species that have been 
vouchered by a single specimen (e.g., Erynnis zarucco) or rarely recorded (e.g., Speyeria 
idalia). It appears to me that these strays should have been included in the main species 
section, which already includes such strays as Pyrgus communis, Hylephila phyleus, 
Battus philenor, to name a few. Next is a section listing five unconfirmed species that 
may range into Ontario based on records from adjacent provinces and Michigan. This 
section might better have been called a hypothetical species list. The final species section 
has information on five doubtful species that have been erroneously referred to Ontario 
as a result of misidentification, improper labelling, or lack of a voucher specimen. 

The last four pages of the Atlas comprise a bibliography, including check-lists, TEA 
publications, and general works, and a provincial ranking of status indicating number of 
occurrences within the province. 

This publication is a valuable addition to the literature on Lepidoptera of the region 
from Hudson Bay to the Great Lakes, although there are a few questionable assertions. 
For example, the authors state that Lycaeides melissa samuelis hibernates “as a larva,” 
but according to James Scott (The Butterflies of North America, 1986, Stanford University 
Press, CA), “eggs hibernate in ssp. samuelis...” And for Danaus plexippus, the authors 
mention “positive evidence for a cycle of abundance peaking about every 11 years.” In 
both cases, the authors fail to include literature citations or to otherwise identify the 
source of this information. 

This Atlas would have been more valuable if specific rearing experiences and personal 
food plant observations had been included. Furthermore, there is no discussion under L. 
m. samuelis or other rare or threatened species as to the authors’ recommendations of 
proposed management methods and techniques to preserve or enhance existing popula- 
tions. This shortcoming appears to be inconsistent with the authors stated purpose “to 
provide a reference for planning efforts to conserve our rare species. ...' Also, I found 
it interesting that the authors assert that the food plants of Erynnis persius are “willows, 
poplars and aspens.” In Michigan, persius feeds on lupine and most recent authoritative 
references state that lupine is the preferred larval food. 

The Ontario Butterfly Atlas, with an 8 x 10” format, is printed on high quality paper 
with easy to read type and excellent photographs. I found no typographical errors or 
improper use of nomenclature. I recommend this publication to all students interested 
in the butterflies and skippers of Ontario and the Great Lakes region. 


MOGENS C. NIELSEN, Adjunct Curator-Lepidoptera, Department of Entomology, 
Michigan State University, East Lansing, Michigan 48824. 


Journal of the Lepidopterists Society 
48(1), 1994, 80-82 


OUT OF AFRICA 


THE BUTTERFLIES OF KENYA AND THEIR NATURAL History, by Torben B. Larsen. 1992. 
Oxford University Press, Oxford, New York and Tokyo. xxii + 490 pp., 19 + A4 figures, 
64 color plates. Hard cover, 18 x 25 cm, ISBN 0-19-854011-6. $195.00. 


BUTTERFLIES OF TANZANIA, by Jan Kielland. 1990. Hill House, Melbourne and London. 
363 pp., 3 maps, 179 text figures, 68 color plates. Hard cover, 22.5 x 30 cm, no ISBN. 
$120.00. 


VOLUME 48, NUMBER 1 81 


These are two rather comparable volumes devoted to the butterfly fauna of adjacent 
East African countries. Both are authoritative, even though Kielland’s volume represents 
many more years of sampling the Tanzanian fauna than does Larsen’s. Despite this 
shortcoming, however, Larsen has managed to produce a very readable and useful volume 
because he has consulted with many authorities and resident Kenyan collectors and has 
organized the field notes of the latter into an excellent compendium. Kielland’s book is 
no less well-documented and both volumes contain much useful information for either 
the collector or the butterfly-watcher. 

The plates, of course, are the highlight of any butterfly book, and these are of top 
quality. The specimens illustrated are the best ones available to the authors: those in the 
Kenya book are presumably the best available at The Natural History Museum, London, 
whereas the specimens in the Tanzanian book are largely those available in Mr. Kielland’s 
collection, sometimes to the detriment of the Tanzanian work. The larger format of 
Kielland’s book makes possible the life-size illustrations of all taxa. Larsen’s book illustrates 
only half of each spread specimen for larger species and has more reduced figures, but 
where applicable, this fact is stated on each caption page. 

The Kenya volume appears to have been proofread somewhat more stringently than 
does the Tanzanian one. A number of annoying “typos” in the latter distract the reader. 
For example, when one seeks the illustration for Bicyclus kiellandi Condamin, the plate 
on which it is figured is stated in the text (page 81) to be Plate 18; the butterfly is actually 
illustrated in Plate 17! Occasional inappropriate capitalizations of some, but not all, 
patronymial specific epithets occur throughout the text. 

Both books use species citations that do not strictly conform to the Rules of Zoological 
Nomenclature because they do not place the authors’ names in parentheses when a species 
or subspecies was described in a genus other than the one in which it is presently contained. 
This habit, while maddening to the purist, seems to be gaining acceptance among authors, 
and perhaps it will no longer be required in future Rules. 

Larsen is a superb writer who manages to use humor to its greatest advantage. He does 
not always write humorously, but rather interjects it occasionally. For example, on page 
34, during a discussion of urine being attractive to butterflies, he provides this aside: 
“(When collecting in the tropics I often take a couple of Johnnie Walker bottles full of 
urine with me; whoever once stole two such bottles from my car must have been in for 
a surprise when he reached home).”’ I suspect that there is more useful information about 
habits of butterflies in the Kenya book, but both provide much useful data. 

I have only two major criticisms, neither perhaps important in itself, but both of which 
are an annoyance. Both books contain a number of descriptions of new taxa, especially 
in the Kielland book, which make the books indispensable to the taxonomist. There seems 
to be a “conspiracy on the part of some authors to include such descriptions in books to 
enhance sales, but perhaps this not intentional. Steve Collins and Larsen do describe their 
new species in an appendix (pp. 438-445) almost apologetically, which seems to mitigate 
my objection mentioned above. Kielland does not so segregate his new names (as well as 
those of other authors), which are included throughout the text. Type specimens are 
illustrated in both books, thereby further increasing their utility to the specialist. 

The second criticism applies only to the Kielland book, and it is not the author’s fault: 
the type faces employed throughout the book are so similar to one another (family-group 
names are in slightly larger type) that it is extremely difficult to locate information quickly 
or determine where one discussion ends and the next begins. In Larsen’s book, the main 
headings are much better accentuated, and one can tell at a glance what constitutes any 
species discussion. In addition, genera are well separated in that text, whereas in the 
Tanzanian volume there is no difference in typography between genera, species or sub- 
species. 

These are really minor complaints, however, and both books are significant additions 
to the African butterfly literature. They are well-written and superbly illustrated, and 
though both volumes are somewhat expensive, they are well worth the money. If the 
reader has to choose only one of the volumes (they do cover roughly the same fauna of 
nearly 900 butterfly species), I would have to recommend Larsen’s volume over Kielland’s 
based chiefly on the clarity of the former’s presentation. Hopefully, no one will have to 


82 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


choose and those interested in east African butterflies can have both volumes—the authors 
are to be congratulated for their accomplishments. 


LEE D. MILLER, Allyn Museum of Entomology of the Florida Museum of Natural 
History, 3621 Bay Shore Road, Sarasota, Florida 34234. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 82-83 


A PRACTICAL GUIDE TO BUTTERFLIES AND MOTHS IN SOUTHERN AFRICA, by S. E. Wood- 
hall (co-ordinating editor) et al. 1992. Lepidopterists’ Society of Southern Africa, P.O. 
Box 470, Florida Hills, 1710, Transvaal, Republic of South Africa [Sponsored by Monsanto 
South Africa (Pyt) Ltd., Agricultural Group]. 223 pp., 48 color photographs on 8 plates, 
numerous black and white photographs and text figures. Softcover, glossy paper, 14.5 x 
21 cm, ISBN 0-620-16774-2. Available from the Lep. Soc. So. Africa for $25 U.S. plus 
postage ($1 for surface mail or $20 for Airmail). 


This techniques guide was compiled through the efforts of nineteen of the top lepi- 
dopterists in southern Africa. It is an easily totable resource (approximately 6” x 8” x 
%2") for almost every aspect of studying Lepidoptera, meant for use by beginners as well 
as by others who desire to learn more about techniques used in other countries. It is easy 
reading, even amusing at times, and the techniques appropriate for southern Africa are 
certainly usable here. Although the price is high for a paperback, the book is absolutely 
stuffed with information! 

Chapter topics include the history of lepidopterists and lepidopterology in southern 
Africa, the lepidopteran life cycle, morphology and terminology, binomial nomenclature, 
species theory, evolution of Lepidoptera, a discussion on “lumpers vs. splitters, ’ taxonomic 
classification, and the rules that govern scientific nomenclature (ICZN). As expected in a 
work meant to address beginners, the book is amply illustrated with black and white 
photographs, line drawings, cartoons, and eight pages of color photographs depicting 48 
live specimens of butterflies, moths (even some micros), and larvae which accompany the 
systematic classification of Lepidoptera and a brief overview of Lepidoptera. 

A chapter devoted to Lepidoptera conservation in southern Africa presents the various 
habitat changes and impacts that man’s activities have had on insect populations and 
includes discussions of monitoring methods (conducting mark-recapture studies, hostplant 
monitoring, habitat monitoring). Also in this chapter is mention of the South African 
Red Data Book—Butterflies, a publication documenting 102 species considered to be 
exposed to some level of threat. This guide provides a table of 190 or so rare, endangered 
or vulnerable butterfly species cross tabulated with habitat, habits, distribution, taxonomy, 
food, and reproduction. Checkmarks in the columns denote a lack of information about 
a specific aspect of the particular butterfly. This table draws attention to those aspects 
unknown, with the idea that special efforts should be made to remedy these “unknowns.” 

Collecting and field techniques for butterflies are detailed and are accompanied by 
photographs, cartoons, and line drawings. Included are instructions on the construction 
of equipment and the proper use of nets, traps, decoys and baits, killing methods, and 
specimen storage in the field, along with suggestions for observing hilltopping, territo- 
riality, and egg laying behavior. Especially interesting are the instructions for preparing 
rotten shrimp bait, various fruit baits, and different combinations of scat and urine to 
attract specimens. Apparently, the fresher the dung, the better (baitwise). However, a 
cautionary note is included to remind the lepidopterist to keep a sharp lookout for the 
originator of the dung; e.g., lions, elephants, baboons, etc. Discussion of moth collecting 
includes traps (many diagrams and photos), sugaring, and the collecting of immatures. 

Preservation and mounting techniques are covered thoroughly. Relaxing techniques 
(chambers, injection, cutting wing muscles), selecting pin sizes, pinning specimens, con- 


VOLUME 48, NUMBER 1 | §3 


struction of various spreading boards, use of various materials as “setting strips,” and 
setting pins are all discussed. The actual spreading technique for butterflies and macros 
is described in detail, with many cautions included for the inexperienced. There are also 
instructions for baking (=drying) specimens while on the spreading board. Although the 
description of techniques for pinning micros is brief, detailed instructions are given for 
preserving and mounting early stages, and for dissection and preservation of genitalia, 
palpi, legs, androconia, and larval head shields. Degreasing, dust removal, mold removal, 
repairing damaged specimens, housing the collection, prevention of infestation, proper 
labelling, and hints for successful mailing of specimens are also included in this chapter. 

Other topics covered in this guide include rearing and breeding Lepidoptera (cages, 
sleeving, foodplants, artificial diets, and larval diseases) and photography in the field and 
in the studio (backdrops and composition, as well as proper storage of photographs). There 
are chapters on good scientific practice, the British Butterfly Conservation Society Code 
of Practice, several appendices (providing addresses of equipment dealers, names and 
addresses of journals worldwide that accept Lepidoptera papers, reviews of threatened 
species in southern Africa), a glossary of terms, a bibliography, and an index. 

Throughout the book the authors encourage all lepidopterists to expand and further 
the science of lepidopterology. Emphasis on observations in the field and the elucidation 
of life histories are recurring themes in this handbook that espouses a common-sense 
approach to the study of Lepidoptera. Amateur lepidopterists are hailed for tirelessly 
providing valuable information, continually taking on the onerous task of breeding and 
recording life histories, and contributing much of what is known about Lepidoptera. The 
debate over collecting vs. no-collecting is broached. Admitting that it is an emotionally 
charged issue, the authors agree that general bans on collecting are detrimental to the 
conservation of Lepidoptera and of nature as a whole. Conservation is promoted and 
responsible collecting is encouraged. The point is made that the reproductive capabilities 
of insects far outweigh the impact of the small numbers collected by lepidopterists. A 
reminder to all is that the younger generation should be encouraged to study invertebrates; 
that first-hand collecting and field experiences, besides being enjoyable, entice children 
to become more aware of nature and more able at a later date to make informed 
judgements on conservation matters; and that if “... no more children are starting at the 
basics, one day there will be no lepidopterists to be consulted on conservation.” 

I found the book to be an inspiring and complete guide to handling Lepidoptera, 
written in an easy to understand, often entertaining manner. It was rewarding to read a 
techniques book that continually encouraged and lauded the efforts of avocational lepi- 
dopterists. It sets an excellent example for our own future Lepidopterists’ Society Tech- 
niques Manual. 


STEPHANIE McKown, 650 Cotterell Drive, Boise, Idaho 83709. 


Journal of the Lepidopterists’ Society 
48(1), 1994, 84 


MANUSCRIPT REVIEWERS, 1993 


The merit of a scientific journal depends on the quality of its reviewers as well as of 
its authors, but the former are usually unknown to readers. The Journal relied on the 
expertise of 68 reviewers last year to provide 89 evaluations of manuscripts. It is with 
much gratitude that the Journal acknowledges the services of the people listed below 
from whom manuscript reviews were received in 1993. 


Phil R. Ackery, London, England 
Annette Aiello, Panama 

Paul H. Arnaud, Jr., San Francisco, CA 
*George T. Austin, Las Vegas, NV 


Susan S. Borkin, Milwaukee, WI 
*M. Deane Bowers, Boulder, CO 


Andrew V. Z. Brower, Ithaca, NY 

Lincoln P. Brower, Gainesville, FL 

*Keith S. Brown, Sao Paulo, Brazil 

*Richard L. Brown, Mississippi State, 
MS 

*John M. Burns, Washington, DC 


John V. Calhoun, Dunedin, FL 

Reginald Chapman, Tempe, AZ 

Frances S. Chew, Medford, MA 

Ian F. B. Common, Toowoomba, 
Queensland, Australia 

Charles V. Covell, Jr., Louisville, KY 


P. T. Dang, Ottawa, Ontario, Canada 

Don R. Davis, Washington, DC 

John J. Dayton, Santa Cruz, CA 

*John A. De Benedictis, Davis, CA 

*Philip J. DeVries, Austin, TX 

Julian P. Donahue, Los Angeles, CA 

*Boyce A. Drummond, Woodland Park, 
CO 


Thomas D. Eichlin, Sacramento, CA 
*John F. Emmel, Hemet, CA 
Marc Epstein, Washington, DC 


David K. Faulkner, San Diego, CA 
Douglas C. Ferguson, Washington, DC 
Clifford D. Ferris, Laramie, WY 


Lawrence F. Gall, New Haven, CT 
*Glenn A. Gorelick, Glendora, CA 


David L. Hancock, Brisbane, 
Queensland, Australia 

John B. Heppner, Gainesville, FL 

Ronald W. Hodges, Washington, DC 


* Reviewed two or more manuscripts. 


Daniel Janzen, Philadelphia, PA 
Kurt Johnson, New York, NY 


Niels P. Kristensen, Copenhagen, 
Denmark 


Gerardo Lamas Muller, Lima, Peru 
Ron Leuschner, Manhattan Beach, CA 


*C. Don MacNeill, San Francisco, CA 
Deborah L. Matthews, Gainesville, FL 
Tim L. McCabe, Albany, NY 
Jacqueline Y. Miller, Sarasota, FL 
William E. Miller, St. Paul, MN 


Raymond W. Neck, Houston, TX 
Mogens C. Nielsen, East Lansing, MI 


Robert W. Poole, Washington, DC 
*Jerry A. Powell, Berkeley, CA 


Frederick H. Rindge, New York, NY 
*Robert K. Robbins, Washington, DC 


Walter H. Sakai, Santa Monica, CA 
David A. Schooley, Reno, NV 
*James A. Scott, Lakewood, CO 
Arthur M. Shapiro, Davis, CA 
Oakley Shields, Mariposa, CA 

John A. Shuey, Traverse City, MI 
Stephen M. Spomer, Lincoln, NE 
Stephen R. Steinhauser, Sarasota, FL 
Ray E. Stanford, Denver, CO 
*Stephen Stone, Lakewood, CO 


Thomas Turner, Clearwater, FL 
Paul M. Tuskes, San Diego, CA 


R. I. Vane-Wright, London, England 
Adrienne Venables, Washington, DC 


*David L. Wagner, Storrs, CT 

Warren H. Wagner, Jr., Ann Arbor, 
MI 

David A. West, Blacksburg, VA 

William D. Winter, Dedham, MA 


Date of Issue (Vol. 48, No. 1): 23 February 1994 


EDITORIAL STAFF OF THE JOURNAL 


JOHN W. Brown, Editor 
Entomology Department 
San Diego Natural History Museum 
POy Box 1390) *- 
San Diego, California 92112 U.S.A. 


Associate Editors: 
M. DEANE BOwERS (USA), BoYCE A. DRUMMOND (USA), LAWRENCE F. GALL (USA), 
GERARDO LAMAS (Peru), ROBERT C. LEDERHOUSE (USA), ROBERT K. ROBBINS (USA), 
CHRISTER WIKLUND (Sweden) 


NOTICE TO CONTRIBUTORS 


Contributions to the Journal may deal with any aspect of Lepidoptera study. Categories 
are Articles, Profiles, General Notes, Technical Comments, Book Reviews, Obituaries, 
Feature Photographs, and Cover Illustrations. Reviews should treat books published within 
the past two years. Obituaries must be authorized by the President of the Society. Re- 
quirements for Feature Photographs and Cover Illustrations are stated on page 111 in 
Volume 44(2). Journal submissions should be sent to the editor at the above address. 
Short manuscripts concerning new state records, current events, and notices should be 
sent to the News, Stephanie McKown, Editor, 650 Cotterell Drive, Boise, Idaho 83709 
U.S.A. For information regarding book reviews, contact Boyce A. Drummond, Natural 
Perspectives, P.O. Box 9061, Woodland Park, CO 80866. Journal contributors should 
submit manuscripts in triplicate, typewritten, entirely double-spaced, with wide margins, 
on one side only of white, letter-sized paper. Prepare manuscripts according to the 
following instructions, and submit them flat, not folded. 

Abstract: An informative abstract should precede the text of Articles and Profiles. 

Additional key words: Up to five key words or terms not in the title should accompany 
Articles, Profiles, General Notes, and Technical Comments. 

Text: Contributors should write with precision, clarity, and economy, and should use 
the active voice and first person whenever appropriate. Titles should be explicit, descrip- 
tive, and as short as possible. The first mention of a plant or animal in the text should 
include the full scientific name with author, and family. Measurements should be given 
in metric units; times in terms of the 24-hour clock (0930 h, not 9:30 AM). Underline 
only where italics are intended. 

Literature Cited: References in the text of Articles, Profiles, General Notes, and 
Technical Comments should be given as Sheppard (1959) or (Sheppard 1959, 196la, 
1961b) and listed alphabetically under the heading LITERATURE CITED, in the following 
format without underlining: 


SHEPPARD, P. M. 1959. Natural selection and heredity. 2nd ed. Hutchinson, London. 
209 pp. 

196la. Some contributions to population genetics resulting from the study of 

the Lepidoptera. Adv. Genet. 10:165-216. 


Illustrations: Only half of symmetrical objects such as adults with wings spread should 
be illustrated, unless whole illustration is crucial. Photographs and drawings should be 
mounted on stiff, white backing, arranged in the desired format, allowing (with particular 
regard to lettering) for reduction to fit a Journal page. Illustrations larger than letter- 
size are not acceptable and should be reduced photographically to that size or smaller. 
The author's name and figure numbers as cited in the text should be printed on the back 
of each illustration. Figures, both line drawings and photographs, should be numbered 
consecutively in Arabic numerals; “plate” should not be employed. Figure legends must 
be typewritten, double-spaced, on a separate sheet (not attached to illustrations), headed 
EXPLANATION OF FIGURES, with a separate paragraph devoted to each page of illustrations. 
Color illustrations are encouraged; contact editor for submission requirements and cost. 

Tables: Tables should be numbered consecutively in Arabic numerals. Headings for 
tables should not be capitalized. Tabular material must be typed on separate sheets, and 
placed following the main text, with the approximate desired position indicated in the 
text. Vertical lines as well as vertical writing should be avoided. 

Voucher specimens: When appropriate, manuscripts must name a public repository 
where specimens documenting identity of organisms can be found. Kinds of reports that 
require vouchering include life histories, host associations, immature morphology, and 
experimental enquiries. 

Proofs: The edited manuscript and galley proofs will be mailed to the author for 
correction of printer's errors. Excessive author’s changes at this time will be charged to 
authors at the rate of $2 per line. A purchase order for reprints will accompany proofs. 

Page charges: For authors affiliated with institutions, page charges are $20 per Jour- 
nal page. For authors without institutional support, page charges are $10 per Journal 
page. Authors unable to pay page charges for any reason should apply to the editor at 
the time of submission for a reduced rate or free publication. Authors of Book Reviews 
and Obituaries are exempt from page charges. 

Correspondence: Address all matters relating to the Journal to the editor. 


PRINTED BY THE ALLEN PRESS, INC., LAWRENCE, KANSAS 66044 U.S.A. 


CONTENTS 


PRESIDENTIAL ADDRESS 19938: ON THE COMPARATIVE DISTRIBU- 
TIONS OF LEPIDOPTERA AND LEPIDOPTERISTS. Ray E. Stan- : 
fot tc ee Reh Sa RS Nb i iin eC eg ] 


A NEW GENUS OF WINTER MOTHS (GEOMETRIDAE) FROM EASTERN 
CALIFORNIA AND WESTERN NEVADA. Jerry A. Powell and 
Douglas C: Ferguson: 22) 600s a 8 


SPLIT SKIPPERS: MEXICAN GENUS POANOPSIS GOES IN THE ORIG- 
ENES GROUP—AND YVRETTA FORMS THE RHESUS GROUP— 
OF POLITES (HESPERIIDAE). John M, Burns ..3) 02a 24 


A NEW SPECIES OF ACROLEPIOPSIS AND THE DESCRIPTION OF THE 
FEMALE OF A. CALIFORNICA (ACROLEPIIDAE). Reinhard 


Gaedike: (00 co RS OO SS A6 
A NEW SPECIES OF LAPARA (SPHINGIDAE) FROM SOUTHEASTERN 
UNITED STATES. Vernon Antoine Brou Jr. 2) ol 
PROFILE 
William Wittfeld: The Florida connection. John V. Calhoun 58 
GENERAL NOTES 
Musineon tenuifolium (Apiaceae): New host of four Nebraska Papilio (Pa- 
pilionidae). “J. M. Reiser and S. M. Spomer, 1... oo) ee 68 
Two new synonymies in Nearctic Eucosma (Tortricidae: Olethreutinae). 
Loran D. Gibson and William E. Miller 69 
Additional data on the geographical distribution and adult activity of the 
diurnal, mimetic plume moth, Oidaematophorus chamelai (Pterophor- 
idae).\J. A. Powell and J. AsChemsak: 2 00 2) 3 ee 71 
Book REVIEWS 
Keys to the insects of the European part of the USSR. Volume IV (Lepidoptera), 
Part T1.. \Joel Mimet fo 2G00h oo Von Ss Oh IY AeA AS a 74 
Butterfly gardening in the south: Cultivating plants that attract butterflies. 
Jacqueline Y. Millere isco s0 285 0 VE 76 
Butterfly gardening: Creating summer magic in your garden. Jacqueline Y. 
Miller: eee yk MN ete CUD SC 78 
The Ontario’ butterfly atlas» “Mogens, C. Nielsen (00... eee 79 
The butterflies of Kenya and their natural history. Butterflies of Tanza- 
nial ee DD. Miller: 3000 Val 80 
A practical guide to butterflies and moths in southern Africa. Stephanie 
ME CK Otay ces S88 aN eh nl UES Saal eee lad ce POEL Fo 
MANUSCRIPT REVIEWERS, 199800 oO 84 


This paper meets the requirements of ANSI/NISO Z39.48-1992 (Permanence of Paper). 


q Volume 48 1994 | Number 2 


oa 


ISSN 0024-0966 


JOURNAL 


of the 


LEPIDOPTERISTS’ SOCIETY 


Published quarterly by THE LEPIDOPTERISTS’ SOCIETY 


Publié par LA SOCIETE DES LEPIDOPTERISTES 
Herausgegeben von DER GESELLSCHAFT DER LEPIDOPTEROLOGEN 
Publicado por LA SOCIEDAD DE LOS LEPIDOPTERISTAS 


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17 May 1994 


THE LEPIDOPTERISTS’ SOCIETY 
EXECUTIVE COUNCIL 


PAUL A. OPLER, President JORGE E. LLORENTE-BOSQUETS, 
Ray E. STANFORD, Immediate Past Vice President 

President FREDERICK W. STEHR, 
CHEN-SHING LIN, Vice President Vice President 
MICHAEL J. SMITH, Secretary ROBERT J. BORTH, Treasurer 
Members at large: 
Charles V. Covell, Jr. Eric H. Metzler John V. Calhoun 
Linda S. Fink Robert K. Robbins Robert C. Lederhouse 
Scott E. Miller J. Benjamin Ziegler William E. Miller 


EDITORIAL BOARD 


PAUL A. OPLER (Chairman), FREDERICK W. STEHR (Member at large) 
JOHN W. BROWN (Journal), WILLIAM E. MILLER (Memoirs) 
STEPHANIE S. MCKOWN (News) 


HONORARY LIFE MEMBERS OF THE SOCIETY 


CHARLES L. REMINGTON (1966), E. G. MUNROE (1978), 
ZDRAVKO LORKOVIC (1980), IAN F. B. COMMON (1987), JOHN G. FRANCLEMONT (1988), 
LINCOLN P. BROWER (1990), DOUGLAS C. FERGUSON (1990), 
HON. MIRIAM ROTHSCHILD (1991), CLAUDE LEMAIRE (1992) 


The object of the Lepidopterists’ Society, which was formed in May 1947 and for- 
mally constituted in December 1950, is “to promote the science of lepidopterology in all 
its branches, .... to issue a periodical and other publications on Lepidoptera, to facilitate 
the exchange of specimens and ideas by both the professional worker and the amateur 
in the field; to secure cooperation in all measures’ directed towards these aims. 

Membership in the Society is open to all persons interested in the study of Lepi- 
doptera. All members receive the Journal and the News of the Lepidopterists’ Society. 
Institutions may subscribe to the Journal but may not become members. Prospective 
members should send to the Treasurer full dues for the current year, together with their 
full name, address, and special lepidopterological interests. In alternate years a list of 
members of the Society is issued, with addresses and special interests. There are four 
numbers in each volume of the Journal, scheduled for February, May, August and 
November, and six numbers of the News each year. 


Active members—annual dues $25.00 
Student members—annual dues $15.00 
Sustaining members—annual dues $35.00 
Life members—single sum $500.00 
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Cover illustration: The buckeye butterfly (Junonia coenia) is a common and widespread 
species of North America. Original drawing by Marianne C. Filbert, P.O. Box 155, Logan, 
Utah 843821. 


JOURNAL OF 


Tue LeEerPIDOPTERISTS’ SOCIETY 


Volume 48 1994 Number 2 


Journal of the Lepidopterists’ Society 
48(2), 1994, 85-105 


SAMPLING STRATEGIES FOR ESTIMATING MOTH 
SPECIES DIVERSITY USING A LIGHT TRAP IN A 
NORTHEASTERN SOFTWOOD FOREST 


A. W. THOMAS 


Canadian Forest Service, Natural Resources Canada, P.O. Box 4000, 
Fredericton, New Brunswick E3B 5P7, Canada 


AND 


G. M. THOMAS 
University of New Brunswick, Fredericton, New Brunswick E8B 6C2, Canada 


ABSTRACT. A 22-watt black-light trap was operated for 29 nights within a forest 
canopy in the Maritime Lowlands Ecoregion of the Acadian Forest. The species-abun- 
dance frequency distribution (pattern of species abundance) was a good fit to the log 
series model and this model was used for subsequent data analysis. No single-night sample 
adequately estimated the log series alpha index of diversity based on the total catch; some 
sampling effort was required each night. Each night’s catch was separated into 16, 30- 
minute samples. The alpha index of diversity for the summed catch for each time-period 
was compared with the overall alpha based on the total catch. A strategy that involved 
operating the trap for just a 1-hour period each night had no effect on the pattern of 
species abundance and gave a value for alpha equal to that obtained by operating the 
trap for an 8-hour period each night. This strategy reduced the catch from 6088 to 971 
moths and the number of species from 255 to 161. Processing costs associated with the 
larger sample and any possible negative effect on the moth population caused by removal 
trapping were greatly reduced. This new sampling strategy is thus useful for comparing 
indices of species diversity between several sites when data are collected simultaneously, 
but is of limited use for species-inventory studies. 


Additional key words: species-abundance distribution, 30-minute samples, log-series 
model, partial-night sampling. 


In recent years, the challenge to maintain biodiversity on this planet 
has become a major public concern. Most attention focuses on Neo- 
tropical ecosystems (Mares 1992). However, the importance of main- 
taining Canada’s biodiversity was addressed in Environment Canada’s 
Green Plan (Hyslop & Brunton 1991), and the launching, in 1991, of 
“Canadian Biodiversity’’ produced by the Canadian Centre of Biodi- 
versity at the Canadian Museum of Nature lends credence to the recent 


86 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


national interest in biodiversity. The values of maintaining current 
biodiversity have been stated by many authors, and summarized by 
Ehrlich (1990) into ethical, aesthetic, economic, and “ecosystem ser- 
vices.’ Salwasser (1990) added the legal obligation for conserving bio- 
logical diversity. Intimately linked with the concept of maintaining 
biodiversity, and especially protection of areas rich in species, is the 
need for a “quick and dirty survey to chart biodiversity of the planet” 
(Roberts 1988), a view reiterated by Ehrlich (1992). The ‘quick and 
dirty’ approach does not advocate poor science; rather, it recognizes 
that the scope of diversity from individual gene systems through pop- 
ulations of species, communities, ecosystems, and ultimately all life in 
the biosphere (Wilson 1988) cannot be addressed in the short-term. It 
suggests that studies should be focused on certain taxonomic groups 
over an extensive area. The hope is that areas with many species or 
high endemism in the selected groups will reflect similarly high values 
for other groups (Roberts 1988). Because of logistic and knowledge 
constraints, the number of species within a community can be deter- 
mined for only a limited number of taxonomic groups. 

This study addresses just one segment of biodiversity, i.e., the diversity 
of moths in a single ecosystem. Diversity is used here to mean the number 
of species and their relative abundance (Magurran 1988), and to prevent 
ambiguity we will always use ‘species diversity’ where appropriate. 
Relative abundance is considered in the form of species-abundance 
frequency distributions, which show the relationship between the abun- 
dance of individuals and the number of species possessing that abun- 
dance (May 1975); abbreviated in this paper as the pattern of species 
abundance. The ecosystem studied is one locality in the Maritime Low- 
lands Ecoregion of the Acadian Forest (Loucks 1962). 

The use of the moth community, in the 15 families used in this study 
(see Appendix), as an exemplar of the species diversity of this ecosystem 
has advantages that include the relative ease of identification at the 
species level, the somewhat standardized sampling methodology (Wil- 
liams 1951, Williams et al. 1955, Taylor & French 1974, Bowden 1982), 
and the high correlation of insects, in general, with the spatial, archi- 
tectural, and taxonomic diversity of plants (Southwood et al. 1979). 

No community consists of species of equal abundance (Magurran 
1988). It is normally the case that the majority of species are rare while 
a number are moderately common with the remaining few species 
being very abundant (Williams 1964, May 1975, Pielou 1975, South- 
wood 1978, Magurran 1988). Within this general distributional form, 
communities have characteristically different patterns of species abun- 
dance which remain stable despite changes in species composition (Pie- 
lou 1975, May 1976, Kempton 1979). The pattern of species abundances 


VOLUME 48, NUMBER 2 87 


at a site allows for comparison with similar sites that have different 
mixes of species, and a change in the pattern of abundance at one site 
has been shown to be a useful indicator of environmental disturbance 
(Kempton & Taylor 1974, Taylor et al. 1978, Kempton 1979). 

Four main species-abundance models (the geometric series, the log- 
arithmic series, the log normal distribution, and MacArthur’s broken 
stick model) have been developed to describe species diversity in terms 
of an ‘index parameter’ as well as the pattern of species abundance. In 
addition there are several non-parametric indices based on the pro- 
portional abundances of species (May 1975, Southwood 1978, Magurran 
1988). 

The log series model was the first to describe the pattern of species 
abundance (Fisher 1943). Since then it has been found to have a wide 
application for catches of many invertebrates, e.g., moths in light traps 
(Williams 1948, 1945, 1964, Taylor & Brown 1972, Taylor & French 
1974, Kempton & Taylor 1974, Taylor et al. 1976, 1978, Taylor 1986), 
Ichneumonidae (Owen & Chanter 1970), cockroaches (Wolda 1983), 
Psocoptera (Broadhead & Wolda 1985), Hymenoptera (Noyes 1989), 
and the community of phytophagous arthropods on apple (Brown & 
Adler 1989). Its wide applicability is because it is based on the abun- 
dances of the species with medium abundance rather than the very 
abundant and very rare species (Taylor et al. 1976, Kempton 1979, 
Brown & Adler 1989). 

The log series is a simple two-parameter model, with two defining 
multispecies population parameters, chi and alpha. Chi is devoted to 
sample characteristics and varies with sample size as it is a function of 
the mean number of individuals per species. Alpha is independent of 
sample size and characterizes the required population quality (Kempton 
& Taylor 1974). Fisher’s (1948) initial suggestion was that alpha might 
‘be useful as a measure of ‘species richness’ when comparing samples. 
Williams (1943) suggested that the parameter alpha be known as a 
community’s ‘index of diversity.’ Later he recognized that this term 
was applicable to other functions having the same properties and re- 
ferred to Fisher’s alpha as ‘diversity calculated on the basis of the 
logarithmic series’ (Williams 1964). The log series model can be derived 
from two statistics, S, the total number of species, and N, the total 
number of moths. It is a discontinuous frequency series with an infinite 
number of terms: 


Me GK 2 Sp IK Aas oe, 
where n, is the number of species with 1 individual and successive 


terms with 2, 3, 4, etc. individuals, and x (chi) is a constant <1 (Williams 
1947). 


88 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


The log normal model was compared with the log series model by 
Kempton and Taylor (1974) in an analysis of moth catches from light 
traps at 18 sites for four successive years in an attempt to quantify 
intuitively recognized properties of habitats. This comparison found 
that samples from stable environments were best fitted by the log series 
whereas those from highly perturbed sites better fitted the log normal. 
Their overall conclusion was that alpha of the log series was the superior 
diversity discriminant, which they defined as a population parameter 
that behaves consistently within a stable population and responds to 
changes within, and to differences between, environments (see also 
Taylor et al. 1976). 

The Simpson-Yule diversity statistic and the Shannon-Weaver infor- 
mation statistic (both non-parametric indices) were compared to the 
log series alpha index of diversity by Taylor et al. (1976) using 10 years 
of light-trap data at one site. Although the log series model was not the 
ideal description of the pattern of species abundance, the site’s envi- 
ronmental stability was better reflected by alpha than by either of the 
other two statistics. 

One constraint with using the moth community as an exemplar of 
species diversity for an ecosystem is the logistics of sorting, counting, 
and identifying all the individuals in the sample (Taylor 1979). For 
example, a one-night catch from one trap in Kenya exceeded 6.7 kg 
(Taylor et al. 1979); 26,300 moths were captured in one light-trap during 
a nine-month period at Rothamsted (UK) (Williams 1964); 118,256 
moths were taken in one light-trap in one year in Kansas (USA) (Wil- 
liams 1945); 6088 moths were taken in one trap in one month (this 
study). Methods for reducing the size of catches were detailed by Taylor 
and Brown (1972), and for subsampling from large catches by Taylor 
et al. (1979). The objectives of this study were: (1) to describe the 
species-abundance frequency distribution and determine the log series 
alpha index of diversity, for moths captured in a light-trap in a within- 
canopy site of a predominantly balsam fir forest during the flight season 
of the major forest pest, spruce budworm (Choristoneura fumiferana 
(Clemens) (Tortricidae)), and (2) to develop a sampling strategy that 
reduced the catch to a minimum without causing significant loss of 
information, measured as no change in the pattern of species abundance 
and a reduction in the alpha value of 5% or less. 


METHODS 


Moth collection and identification. Beginning on 21 June 1990 (day 
1) and ending on 30 July (day 40), one 22-watt black-light trap (Uni- 
versal Light Trap, Bioquip Products, California) was operated in the 
Peter Brook study area of the Acadia Forest Experiment Station near 


VOLUME 48, NUMBER 2 89 


Fredericton, New Brunswick, Canada. For a variety of reasons, full- 
night trap data are available for only 29 of the potential 40 nights. 
Intensive studies on the population dynamics of spruce budworm have 
been in progress at this site since 1986. The physical characteristics and 
vegetation of the site have been described (Lethiecq & Regniére 1988). 
Briefly, the study area is composed of 77% balsam fir, Abies balsamea 
(L.) Miller (Pinaceae), 12% red maple, Acer rubrum L. (Aceraceae), 
and eight other tree species. However, the surrounding area is heter- 
ogenous and within a 10-km radius contains mixed forest, lakes, streams, 
sphagnum bogs, large clear-cuts, and roadsides. 

The trap, with the lamp at 6.4 m above the ground, was on a platform, 
3 x 1.5m, on a tower within the closed crowns of balsam fir trees; the 
otherwise touching branches were trimmed to leave a clearing of 3 x 
1.5 m. A blue plastic sheet, 1.8 x 2.4 m, was stretched above the platform 
at a height of 2.4 m above the lamp. This sheet made direct observation 
of the light impossible from above, although the reflection of the light 
off of the foliage of the adjacent trees gave a glow to the immediate 
area which was obvious from the ground. 

The trap was equipped with an automatic time-interval collecting 
device (King et al. 1965, Smith et al. 1973). Each night’s total catch 
consisted of 16, 30-minute sequential samples, beginning with time- 
period | from 2130-2200 h and ending with time-period 16 from 0500- 
0530 h. On 21 June, day 1, sunset was at 2120 h and sunrise the following 
morning at 0536 h; on 30 July, day 29, sunset was at 2058 h and sunrise 
the following morning at 0606 h. At the latitude of New Brunswick, 
the sky is noticeably lighter about 30 min before sunrise and remains 
light for 30 min after sunset. 

The moths were killed with 1,1,1 trichloroethane. Moths were stored 
at —17°C until identified and counted. Most specimens were identified 
with the aid of the literature and confirmed by consulting the Forest 
Insect and Disease Survey (FIDS) Reference Collection, Canadian For- 
est Service, Fredericton, which contains specimens identified by the 
Biological Resources Division (BRD) of the Centre for Land and Bio- 
logical Resources Research, Ottawa. Genitalia mounts of specimens 
were made when identification was uncertain. A further 52 species of 
geometrids were identified by Klaus Bolte and 81 species of noctuids 
by Don Lafontaine, both at BRD. All moths in the following families 
were identified to species and counted: Hepialidae; Sesiidae; Cossidae; 
Limacodidae; Thyatiridae; Drepanidae; Geometridae, except for Eu- 
pithecia; Lasiocampidae; Saturniidae; Sphingidae; Notodontidae; Arc- 
tiidae; Lymantriidae; and Noctuidae. In addition, all specimens of spruce 
budworm (Tortricidae) were counted. Moths belonging to other families 
were not identified or recorded. Publications used for species identi- 


90 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


fication were Forbes (1954), McGuffin (1967, 1972, 1977, 1981), Rock- 
burne and Lafontaine (1976), Ferguson (1978), Morris (1980), McCabe 
(1980), Covell (1984), Laplante (1985), Lafontaine (1987), and Lafon- 
taine and Poole (1991). 

Species-abundance frequency distribution. The numbers of species 
having abundances of 1, 2, 3, .. ., 724 moths (based on the total catch) 
were compared with the expected numbers from the log series model 
(Williams 1947) for goodness-of-fit, using the chi-square test (Owen & 
Chanter 1970, Kempton & Taylor 1974, Taylor et al. 1976, Broadhead 
& Wolda 1985, Magurran 1988, Noyes 1989, Basset & Kitching 1991). 
The observed abundances covered a large range, 1-724 moths per 
species, and because many of these 724 abundance classes were zero 
(e.g., abundance classes 31 and 36 each had two species, but no species 
had just 32, 33, 34, or 35 moths and thus classes 832-85 were zeros) the 
abundance classes were grouped into 10 new abundance classes of 
approximately equal range on the logarithmic (base 2) scale (Kempton 
& Taylor 1974, Kempton 1975, Taylor et al. 1976). Because the abun- 
dance class having >511 moths had an expected frequency of <1 
species, this class was pooled with the preceding class to give an expected 
frequency of >1 species; resulting in just nine abundance classes. This 
grouping and pooling of abundance classes (see Table 2) resulted in the 
data set meeting the requirements for the chi-square analysis in that 
no more than 20% of the classes had an expected frequency of <5 
species (1 out of 9 did) and no expected frequency was <1 (Zar 1984). 

Index of diversity. For the purpose of this study, the 29-night sample 
from the trap was taken to be the population being sampled. The log 
series alpha index of diversity was determined after rearranging equa- 
tions (7) and (8) of Williams (1947) to obtain: 


[1] (Sx/—In(’— x)) — Nd —x) —0 
and solving for x using MathCad (1991), and then solving [2] for alpha: 
[2] alpha = N(1 — x)/x 


This value based on the single 29-night sample was termed ‘the overall 
alpha.’ 

Strategies to reduce sample size. Three data manipulations were 
employed to determine a strategy that would reduce the size of the 
sample and thus reduce processing costs and lessen the possible effect 
of removal trapping on the moth population. 

Single-night samples. The first attempt at a sampling strategy was 
to determine alpha for each night’s catch and to compare each value 
with the overall alpha. Such a strategy would certainly reduce sample 
size, but it was not known how representative such an alpha based on 


VOLUME 48, NUMBER 2 91 


one night’s catch would be of the overall alpha based on the total 29- 
day catch. 

‘Replicated’ single-time-period samples. As each night’s catch con- 
sisted of 16 sequential 30-minute samples, there were 16 single-time- 
period samples, with each sample ‘replicated’ for 29 nights. The alpha 
index of diversity was calculated for each pooled time period (e.g., all 
the moths trapped during time period 1 were pooled) and compared 
with the overall alpha. If an index equivalent to the overall index could 
be estimated from a single 30-minute sample taken each night for 29 
nights, significant saving in processing costs would occur, i.e., 1 x 29 
_ = 29 samples instead of 16 x 29 = 464. 

Truncated samples. This strategy was based on the results of the 
single-time-period analysis. As certain time periods gave low alpha 
values, it was argued that these time periods could be eliminated (thus 
reducing the number of samples, the number of moths, the processing 
costs) without significant loss of information. Two sub-strategies were 
employed. The first, termed early truncation, was to discard cumulative 
sequential time periods from the entire data set beginning with all 29 
samples from time period 1, then all 58 samples from time period 1 + 
time period 2, etc. After 15 truncations only the data set from time 
period 16 remained. The alpha index of diversity was calculated from 
the data set remaining after each truncation and compared with the 
overall alpha to determine the percentage change. Also after each 
truncation, the pattern of species abundance was compared with that 
from the log series model using the deviance chi-square values (Kemp- 
ton & Taylor 1974). The second sub-strategy, termed late truncation, 
was similar to early truncation except that all 29 samples from time 
period 16 were first discarded, then all 58 samples from time periods 
16 + 15, etc. Combining selected data sets that remained after early 
and late truncation (effectively a double-ended truncation) gave several 
sampling strategies that met the goal of reducing sample size without 
compromising the value for alpha or the pattern of species abundance. 
The durations for these sampling strategies are shown in Table 1. 


RESULTS 


Totals of 6088 individual moths representing 255 macrolepidoptera 
species in 15 families were identified from the 29-night catch (see 
Appendix). 

Species-abundance distribution and index of diversity. The pattern 
of species abundance is shown in Table 2. In general, the number of 
species in the abundance classes decreased as the abundance increased. 
Most species (52) were in the first abundance class, making this the 


92 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


TABLE 1. Time-periods for sampling strategies. 


Strategy # Inclusive time-periods Extent of sample (h) 
] 1-16 2130-0530 
2 3-10 2230-0230 
3 3-9 2230-0200 
4 4-10 2300-0230 
3) 4-9 2300-0200 
6 5-10 2330-0230 
a 5-9 2330-0200 
8 6-10 2400-0230 
9 6-9 2400-0200 

10 7-10 0030-0230 
Ia 7-9 0030-0200 
12 8-10 0100-0230 
13 8-9 0100-0200 
14 9-10 0130-0230 


commonest class. The apparent paradox is that members of these species 
were rare with just one moth in each species (see Appendix). The fewest 
species (8) were in the largest abundance class making this the rarest 
class but members of these species were abundant (>255 moths in each, 
see Appendix). Also shown in Table 2 are the frequencies expected 
from the log series model. The similarity between observed and ex- 
pected appears close and is confirmed as being a good fit by the deviance 
chi-square value of 8.6. The 5% critical value of the chi-square distri- 
bution with 7 df is 14.1 indicating that the log series model provides a 
good description of the data. The overall alpha index of diversity was 54. 

Single-night samples. The number of moth species and individuals 
trapped in a single night ranged from a low value of 30 moths in 18 
species to a high value of 548 moths in 88 species. Values for alpha 


TABLE 2. Species abundance frequency distribution of a moth catch in the Acadia 
Forest Experiment Station compared with expected frequencies from the log series model. 


Number of species 
Individuals 


per species Observed Expected Chi-square 
] 52 53.4 0.04 
2-3 47 43.9 0.22 
4-7 42 39 0.23 
8-15 36 30.4 0.01 
16-31 37 31.3 1.04 
32-63 22 25.3 0.43 
64-127 a 16.9 5.80 
128-255 9 7.8 0.18 

256-511 2 1.76 

On 5, fs aie bier 0.68 


Total chi-square = 8.6, P < 0.5, P > 0.1, df = 7. Last abundance class pooled with previous class to meet requirements 
of chi-square test (see Methods). 


VOLUME 48, NUMBER 2 93 


@ 
fe) 
tobi 
oO 
oO 


30 - 


ALPHA AS PERCENTAGE OF TOTAL 
a 
° 
o 
o 
o 


20 Miner nero tal acl ae a eee 
2 4 6 8 10 12 14 16 18 20 22 24 26 28 
SAMPLE NIGHT 


Fic. 1. Alpha index of diversity for single-night catches as a percentage of the overall 
alpha based on the total catch. 


fluctuated wildly between 12 and 40 with no meaningful trend and 
never closely approaching the overall alpha (Fig. 1). It was apparent 
that no single-night sample could be used to estimate the index of 
diversity and thus no pattern of species abundance was determined. 
‘Replicated’ single-time-period samples. For any single ‘replicated’ 
time period (consisting of 29, 30-minute samples) the total number of 
moths trapped ranged between 48 and 627 and the total number of 


110 


a 
o oO 
om ©) 
Le 
O 
O 
O 
O 


40 


ALPHA AS PERCENTAGE OF TOTAL 
N 
fe) 


ae) [eee [ae eos mL al ca Pee eR 
1 2 3 4 5 6 7 8 989 10 11 12 13 14 15 16 
TIME PERIOD 
Fic. 2. Alpha index of diversity for single-time-period catches, averaged over 29 


nights, as a percentage of overall alpha based on the total catch. Time periods are 
sequential 30-minute periods starting at 2130-2200 h and ending at 0500-0530 h. 


94 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


PERCENTAGE CHANGE IN ALPHA 
nN 
a 
> 
o 


O 
2) a a a a a ei | | 
2 3 4 5 6 7 8 8 10 11 12 183 14 15 16 

TIME PERIOD AT START OF TRAPPING 


PERCENTAGE CHANGE IN ALPHA 
o 
Oo 


q70 Ee eae 
1 2 3 4 5 6 7 8, © 10-141 : 1231325145515 
TIME PERIOD AT END OF TRAPPING 
Fic. 3. Percentage change in the alpha index of diversity compared to the overall 
alpha: (A) when trapping starts with successively later time periods and ends with time 
period 16 (0500-0530 h); (B) when trapping starts at time period 1 (2130-2200 h) and 
end at successively later time periods. 


species trapped ranged between 28 and 182. The values for alpha for 
the ‘replicated’ single-time-period samples started low in the first part 
of the night, rose rapidly to a maximum during the middle part of the 
night and then decreased towards dawn (Fig. 2). For time period 5 the 
value for alpha was 102% that of the overall alpha. However, this datum 
was an outlier that did not follow the trend and it was not thought 


VOLUME 48, NUMBER 2 95 


7000 4 
6000 = o 
5000 a) 
4000 — 


3000 —| 


NUMBER OF MOTHS # 
a 
a 


2000 = 


PERCENTAGE CHANGE IN ALPHA 0 


123 4 5 6 7 8 98 10 11 12 13 14 
SAMPLING STRATEGY 


Fic. 4. Percentage change (empty rectangles) in the alpha index of diversity relative 
to the overall alpha (strategy 1), and number of moths trapped (solid rectangles) for the 
various sampling strategies. See methods and Table 1 for explanation of sampling strat- 
egies. 


prudent to accept this single time period as representative of the overall 
alpha. 

Truncated samples—early truncation. Discarding the data in time 
periods 1 through 6 had no significant effect on alpha determined from 
the remaining data set (Fig. 8A). That is, if the light trap had begun 
operating at 0030 h, start of period 7, and had run until 0530, alpha 
would have been within 5% of the value obtained by starting the light 
trap at 2180 h. Also, early truncation of time periods 1 through 6 had 
no effect on the pattern of species abundance in the remaining data 
set (time-periods 7-16), chi-square = 10.2, 7 df (P > 0.1). 

Late truncation. Discarding the data in time periods 16 through 9 
had no significant effect on alpha based on the remaining data set (Fig. 
3B). That is, if the light trap had begun operating at 2130 h and had 
run until 0130 h, the end of period 8, alpha would have been within 
5% of the overall alpha. Also, late truncation had no effect on the 
pattern of species abundance in the remaining data set (time periods 
1-8), chi-square = 7.7, 6 df (P > 0.1). 

Double-ended truncation. Several combinations of early- and late- 
truncation provided 13 sampling strategies that reduced the sampling 
period and reduced the number of moths trapped. These strategies 
(Table 1) had no significant effect on alpha and did not compromise 
the pattern of species abundance. No calculated chi-square value, com- 
parison between observed pattern of species abundance and expected 


96 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


pattern from the log series model, was significant (P > 0.05). When 
the sampling strategies were arranged in a sequence of decreasing 
sampling periods (Fig. 4), the downward trend in the number of moths 
trapped and the insignificant effect on alpha became obvious. The most 
cost-effective strategy was a 1-h sample obtained nightly from 0130- 
0230 h (strategy 14, Fig. 4) that resulted in a total sample of 971 moths 
in 161 species giving an alpha value of 55. 


DISCUSSION 


The inadequacy of a single-night sample to estimate accurately the 
alpha index of diversity for moths caught during a one-month period 
was observed by Williams (1948, 1964) in England. Nightly samples 
during the month of July gave alpha values that varied from 42-81% 
of the overall alpha based on the total catch for the whole month, with 
no evidence of any regular trend (Williams 1964, Table 67). Taylor 
and Brown (1972) presented data from two traps for nine days in July 
in Kenya. Single-night alpha values ranged from 30.5-80% of the two 
overall values. Our data showed a similar random pattern with nightly 
values varying from 22-74% of the overall alpha value. Even when 
Williams (1964, Table 67) calculated diversity on a weekly basis, the 
average weekly value for alpha was only 77% of the monthly value. 
These data support our conclusion that some sampling effort is required 
nightly throughout the duration of the calendar dates of interest. 

Taylor (1979) commented on the cost-efficiency of sampling insects 
and the advantages of an attractant trap, such as a light trap, in selecting 
specific taxa. He also noted that, when used to control pest-species, light 
traps have as an objective the removal of as large a proportion of the 
population as possible. However, when used as a monitoring tool, the 
objective is to affect the population as little as possible compatible with 
obtaining adequate numbers for analysis. As mentioned in the intro- 
duction, large samples have problems associated with the cost of sorting, 
identifying, counting, and data handling. Reducing sample size by 
subsampling from a larger sample has drawbacks (Taylor et al. 1979). 
Taylor and Brown (1972) tried several methods to decrease the size of 
the moth catch in light traps that included obscuring the light with 
black paint, changing the source of illumination (different bulb types), 
and changing the direction of illumination. These methods reduced the 
size of the catch, but had no effect on the alpha index of diversity. 
They did not examine the effect on the pattern of species abundance. 

Our technique of a short-time-period ‘replicated’ nightly sample to 
determine the alpha index of diversity without changing the pattern 
of species-abundance is new. Because it results in a relatively small 
sample, it has the advantage of affecting the moth population much 


VOLUME 48, NUMBER 2 97 


less than a full-night sample. It appears to be of use for determining 
the moth species diversity of several sites simultaneously which oth- 
erwise could not be considered because of processing costs associated 
with the usually large catches in light traps. 

There are no alpha index of diversity values from eastern North 
American forests in similar latitudes with which to compare the alpha 
value obtained in this study. The long-recognized latitudinal and lon- 
gitudinal gradients in species diversity (Pianka 1966, Smith 1980, see 
also refs. in Magurran 1988) preclude comparison of the alpha value 
from this study with alpha values for moth species diversity in two 
mid-west American states (Williams 1945) and England (Taylor et al. 
1978). 


ACKNOWLEDGMENTS 


We thank Georgette Smith for allowing access to the FIDS reference collection and 
helping with identification. We are especially grateful to Don Lafontaine and Klaus Bolte 
for their skill in the identification of many specimens that were in very poor condition. 


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Received for publication 10 June 1993; revised and accepted 9 October 1993. 


100 


APPENDIX. Species list with numbers of moths and extreme dates of capture. 


Hepialidae 

Korscheltellus gracilis (Grt.) 
Sesiidae 

Synanthedon acerni (Clem. ) 


Cossidae 
Prionoxystus macmurtrei (Guer.) 


Tortricidae 
Choristoneura fumiferana (Clem.) 


Limacodidae 


Tortricidia testacea Pack. 
Tortricidia flexuosa (Grt.) 
Packardia geminata (Pack.) 
Lithacodes fasciola (H.-S.) 


Thyatiridae 
Habrosyne scripta (Gosse) 


Drepanidae 


Drepana arcuata Wk. 
Drepana bilineata (Pack.) 
Oreta rosea (WIk.) 


Geometridae 


Protitame virginalis (Hulst) 
Itame pustularia (Gn.) 

Itame brunneata (Thunb.) 
Itame anataria (Swett) 
Semiothisa aemulataria (Wlk.) 
Semiothisa ulsterata (Pears. ) 
Semiothisa transitaria (Wlk.) 
Semiothisa minorata (Pack.) 
Semiothisa bicolorata (F.) 
Semiothisa bisignata (Wlk.) 
Semiothisa sexmaculata (Pack.) 


Semiothisa signaria dispuncta (Wlk.) 


Semiothisa pinistrobata Fgn. 
Semiothisa orillata (Wlk.) 
Iridopsis larvaria (Gn.) 

Ectropis crepuscularia (D. & S.) 
Protoboarmia porcelaria (Gn.) 
Melanolophia canadaria (Gn.) 
Eufidonia convergaria (Wlk.) 
Biston betularia cognataria (Gn.) 
Hypagyrtis piniata (Pack.) 
Lomographa vestaliata (Gn.) 
Cabera erythemaria Gn. 
Cabera variolaria Gn. 
Euchlaena obtusaria (Hbn.) 
Euchlaena johnsonaria (Fitch) 
Euchlaena marginaria (Minot) 
Euchlaena tigrinaria (Gn.) 
Euchlaena irraria (B. & McD.) 


23-24 July 
25 June-24 July 
28 June 

2-29 July 


26 June-4 July 

25 June-29 July 
21 June-29 July 
26 June-21 July 


25 June-18 July 


21 June-21 July 
26 June-29 July 
10-29 July 


21 June-22 July 
14-29 July 
25 June-17 July 
17 July 
19 July 
29 June 
18 July 
21 June-29 July 
16-19 July 
29 June-19 July 
27 June—24 July 
21 June-29 July 
25 June-25 July 
25-28 June 
21 June-17 July 
27 June-29 July 
25 June-24 July 
21-26 June 
25 June-20 July 
27 June—25 July 
26 June-29 July 
21 June-15 July 
21 June-25 July 
21 June-24 July 
17-18 July 
15-24 July 
25 June 

4-18 July 

2 July 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


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VOLUME 48, NUMBER 2 


APPENDIX. 


Xanthotype urticaria Swett 
Pero morrisonaria (Hy. Edw.) 
Nacophora quernaria (J. E. Smith) 
Campaea perlata (Gn.) 
Tacparia atropunctata (Pack.) 
Tacparia detersata (Gn.) 
Homochlodes fritillaria (Gn.) 
Metanema inatomaria Gn. 
Metanema determinata Wk. 
Metarranthis amyrisaria (W1k.) 
Metarranthis hypocharia (H.-S.) 
Anagoga occiduaria (Wlk.) 
Probole amicaria (H.-S.) 
Plagodis serinaria H.-S. 
Plagodis phlogosaria (Gn.) 
Plagodis alcoolaria (Gn.) 
Caripeta divisata Wlk. 

Caripeta piniata (Pack.) 
Caripeta angustiorata Wk. 
Besma endropiaria (G. & R.) 
Sicya macularia (Harr.) 
Eusarca confusaria Hbn. 
Tetracis cachexiata Gn. 
Nematocampa resistaria (H.-S.) 
Nemoria mimosaria (Gn.) 
Cyclophora pendulinaria (Gn.) 
Scopula cacuminaria (Morr.) 
Scopula limboundata (Haw. ) 
Dysstroma citrata (L.) 
Dysstroma walkerata (Pears.) 
Dysstroma hersiliata (Gn.) 
Eulithis explanata (Wlk.) 
Ecliptopera silaceata albolineata (Pack.) 
Hydriomena perfracta Swett 
Hydriomena renunciata (Wlk.) 
Hydria undulata (L.) 
Rheumaptera hastata (L.) 
Rheumaptera subhastata (Nolcken) 
Mesoleuca ruficillata (Gn.) 
Spargania magnoliata Gn. 
Perizoma basaliata (Wlk.) 
Xanthorhoe abrasaria congregata (W\k.) 
Xanthorhoe iduata (Gn.) 
Xanthorhoe ferrugata (Cl.) 
Xanthorhoe lacustrata (Gn.) 
Hydrelia lucata (Gn.) 

Hydrelia inornata (Hulst) 
Eubaphe mendica (Wlk.) 
Horisme intestinata (Gn.) 
Lobophora nivigerata Wk. 


Lasiocampidae 


Malacosoma disstria Hbn. 
Malacosoma americanum (F.) 


Continued. 


25 June-21 July 
21 June-4 July 
26 June-18 July 
21 June-25 July 
27 June 

21-26 June 
21-29 June 

21 June-29 July 
18-22 July 
21-28 June 

25 June 

21 June 

21 June-9 July 
27 June 

26-29 June 

21 June-4 July 
25 June-29 July 
21 June-23 July 
17-24 July 
21-29 June 
16-25 July 

16 July 

21 June-2 July 
17-29 July 
14-15 July 

21 June-25 July 
18 July 

25 June—24 July 
25-28 June 

21 June-14 July 
15-29 July 
16-29 July 

21 June 

21-25 June 

21 June-29 July 
14 July 

16 July 

26 June-8 July 
25 June 

14 July 

25 July 

25 June-13 July 
12 July 

21 June-4 July 
16 July 

26 June-18 July 
25 June-17 July 
17-20 July 

29 June 

26 June-29 July 


9-29 July 
15-25 July 


101 


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102 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


APPENDIX. Continued. 


Saturniidae 
Dryocampa rubicunda (F.) 21 June-21 July 31 
Anisota virginiensis (Drury) 25 June 1 
Antheraea polyphemus (Cram.) 21 June-22 July 8 
Sphingidae 
Ceratomia undulosa (Wlk.) 21 June 2 
Sphinx gordius Cram. 21 June-20 July 9 
Lapara bombycoides Wk. 21 June-24 July 18 
Smerinthus jamaicensis (Drury) 21 June-25 July 14 
Smerinthus cerisyi Kby. 21-29 June 2 
Paonias excaecatus (J. E. Smith) 21 June-23 July 15) 
Pachysphinx modesta (Harr.) 21 June-23 July 43 
Notodontidae 
Clostera apicalis (Wlk.) 21-26 June 2 
Nadata gibbosa (J. E. Smith) 21 June—-24 July 16 
Peridea basitriens (WIlk.) 15-29 July 2 
Peridea angulosa (J. E. Smith) 24-25 July 2 
Peridea ferruginea (Pack.) 26 June-25 July 150 
Pheosia rimosa Pack. 27 June-29 July 8 
Odontosia elegans (Stkr.) 17-25 July 2 
Notodonta simplaria Graef 15-24 July Ul 
Gluphisia septentrionis W1k. 25 June-25 July 54 
Furcula cinerea (Wlk.) 29 June-24 July 5 
Furcula modesta (Hudson) 16-25 July 1) 
Symmerista leucitys Franc. 21 June 2 
Macrurocampa marthesia (Cram.) 15-25 July 3 
Heterocampa umbrata Wk. 25 June-4 July ll 
Heterocampa guttivitta (Wlk.) 29 June 1 
Heterocampa biundata Wk. 21 June-20 July J4e 
Lochmaeus manteo Doubleday 20-25 July 3 
Schizura ipomoeae Doubleday 21 June-24 July 29 
Schizura badia (Pack.) 21-27 June 2 
Schizura unicornis (J. E. Smith) 15-24 July 10 
Schizura leptinoides (Grt.) 25 June—23 July 8 
Oligocentria semirufescens (Wlk.) 18-24 July 8 
Oligocentra lignicolor (W1k.) 26 June-29 July 89 
Arctiidae 
Eilema bicolor (Grt.) 12-25 July 22 
Hypoprepia fucosa Hbn. 4-29 July 54 
Haploa lecontei (Guer.-Meneville) 9 July 1 
Holomelina laeta (Guer.-Meneville) 29 June—25 July 31 
Holomelina aurantiaca (Hbn.) 20 July il 
Holomelina ferruginosa (Wlk.) 11-22 July Ti 
Pyrrharctia isabella (J. E. Smith) 14 July if 
Spilosoma congrua Wk. 21 June-5 July 40 
Spilosoma virginica (F.) 21 June-20 July 39 
Hyphantria cunea (Drury) 21 June-24 July 182 
Platarctia parthenos (Harr.) 27 June-17 July 8 
Apantesis virguncula (W. Kby.) 27 June-20 July 5 
Halysidota tessellaris (J. E. Smith) 13-19 July 2 
Lophocampa maculata Harr. 21-29 June 48 


VOLUME 48, NUMBER 2 


APPENDIX. Continued. 


Cycnia tenera Hbn. 
Ctenucha virginica (Esp.) 


Lymantriidae 


Dasychira plagiata (Wlk.) 
Leucoma salicis (L.) 


Noctuidae 


Idia americalis (Gn.) 

Idia aemula Hbn. 

Idia rotundalis (Wlk.) 
Zanclognatha pedipilalis (Gn.) 
Zanclognatha protumnusalis (Wlk.) 
Zanclognatha cruralis (Gn.) 
Palthis angulalis (Hbn.) 
Bomolocha baltimoralis (Gn.) 
Lomanaltes eductalis (W1k.) 
Spargaloma sexpunctata Grt. 
Pangrapta decoralis Hbn. 
Parallelia bistriaris Hbn. 
Catocala sordida Git. 
Chrysanympha formosa (Grt.) 
Autographa precationis (Gn.) 
Autographa mappa (G. & R.) 
Syngrapha altera (Ottol.) 
Syngrapha octoscripta (Grt.) 
Syngrapha epigaea (Grt.) 
Syngrapha viridisigma (Grt.) 
Syngrapha alias (Ottol.) 
Syngrapha cryptica Eichlin & Cunningham 
Syngrapha rectangula (W. Kby.) 
Syngrapha microgamma nearctica F gn. 
Plusia venusta Wk. 

Baileya ophthalmica (Gn.) 
Lithacodia muscosula (Gn.) 
Lithacodia synochitis (G. & R.) 
Lithacodia concinnimacula (Gn.) 
Lithacodia carneola (Gn.) 
Leuconycta diphteroides (Gn.) 
Panthea acronyctoides (W\k.) 
Panthea pallescens McD. 
Charadra deridens (Gn.) 

Raphia frater Grt. 

Acronicta americana (Harr.) 
Acronicta dactylina Gtrt. 
Acronicta lepusculina Gn. 
Acronicta innotata Gn.- 
Acronicta tritona (Hbn.) 
Acronicta grisea Wlk. 

Acronicta superans Gn. 
Acronicta hasta Gn. 

Acronicta fragilis (Gn.) 
Acronicta clarescens Gn. 
Acronicta retardata (Wlk.) 


26 June 
10-19 July 


26 June-25 July 
4-19 July 


21 June-29 July 
14-20 July 
21 July 
18-24 July 
12-22 July 
21 June-29 July 
8-29 July 
26 June—20 July 
25 June 
21 June-21 July 
21 June-20 July 
21 June-29 July 
24-25 July 
9-12 July 
29 June 
26 June 
26 June-21 July 
14 July 
15-21 July 
18-24 July 
21 June-20 July 
24 July 
6-25 July 
21 June 
17-19 July 
21 June 
21 June-10 July 
8 July 
25 June—4 July 
25 June—20 July 
21 June-19 July 
21 June-25 July 
27 June-25 July 
21 June-15 July 
21 June-29 July 
21 June-24 July 
14-25 July 
29 June—4 July 
21 June—25 July 
15-19 July 
21 June-24 July 
15 July 
25 June 
21 June—25 July 
25 June-25 July 
26 June—25 July 


103 


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TE RENE TENN NE BRE ENWUDOHENODAIOH 


104 


APPENDIX. 


Acronicta impleta Wk. 
Acronicta noctivaga Grt. 
Acronicta impressa Wk. 


Acronicta oblinita (J. E. Smith) 


Agriopodes fallax (H.-S.) 


Harrisimemna trisignata (Wlk.) 


Apamea verbascoides (Gn.) 
Agroperina cogitata (Sm.) 
Amphipoea velata (W\k.) 
Euplexia benesimilis McD. 
Phlogophora iris Gn. 
Chytonix palliatricula (Gn.) 
Dypterygia rozmani Berio 
Hyppa xylinoides (Gn.) 
Nedra ramosula (Gn.) 
Callopistria mollissima (Gn.) 
Callopistria cordata (Ljungh) 
Proxenus miranda (Gtt.) 
Elaphria versicolor (Grt.) 
Elaphria festivoides (Gn.) 
Apharetra purpurea McD. 
Oncocnemis riparia Morr. 
Polia nimbosa (Gn.) 

Polia imbrifera (Gn.) 

Polia purpurissata (Grt.) 
Polia detracta (Wlk.) 

Polia goodelli (Grt.) 

Polia latex (Gn.) 

Melanchra adjuncta (Gn.) 
Melanchra assimilis (Morr.) 
Lacanobia subjuncta (G. & R.) 
Lacanobia grandis (Gn.) 
Lacanobia lutra (Gn.) 
Lacanobia rugosa (Morr.) 
Lacanobia legitima (Gtrt.) 
Papestra biren (Goeze) 
Lacinipolia lustralis (Grt.) 
Lacinipolia anguina (Grt.) 
Lacinipolia renigera (Steph. ) 
Lacinipolia lorea (Gn.) 
Lacinipolia olivacea (Morr.) 
Leucania multilinea W\k. 
Leucania insueta Gn. 
Leucania inermis (Fbs.) 
Leucania pseudargyria Gn. 
Homorthodes furfurata (Grt.) 
Orthodes crenulata (Butler) 
Orthodes cynica Gn. 

Euxoa divergens (Wlk.) 
Ochropleura plecta (L.) 
Diarsia jucunda (W\k.) 
Eurois astricta Morr. 

Xestia dolosa Franc. 


Xestia oblata (Morr.) 


Continued. 


29 June 
27-28 June 
26 June 
26 June-14 July 
25 June-29 July 
15-29 July 
23 July 
10 July 
23-25 July 
21 June-25 July 
27 June—25 July 
21 June-25 July 
25 June 
18-21 July 
24 July 
25 June-25 July 
21 June-29 July 
4 July 
21 June-13 July 
25 June-20 July 
15-29 July 
14 July 
24-25 July 
18-25 July 
24-25 July 
4-20 July 
16 July 
21 June-10 July 
21 June-24 July 
26 June-25 July 
24 July 
21-28 June 
21 June-25 July 
27 June-16 July 
29 June—25 July 
27 June 
27 June-24 July 
27 June 
29 July 
26 June-18 July 
29 July 
9-25 July 
26 June-21 July 
9-16 July 
4 July 
2-24 July 
25 June-25 July 
21 June-29 July 
5-19 July 
21 June-25 July 
13-29 July 
24-29 July 
15-29 July 
8-20 July 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


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VOLUME 48, NUMBER 2 105 


APPENDIX. Continued. 


Anomogyna elimata (Gn.) 25-29 July 2 
Anomogyna badicollis (Grt.) 24-29 July 5 
Anomogyna youngii (Sm.) 15 July 1 
Aplectoides condita (Gn.) 21 June-4 July 25 
Anaplectoides prasina (D. & S.) 8-29 July 8 
Anaplectoides pressus (Grt.) 15-25 July 4 
Eueretagrotis perattenta (Grt.) 17-25 July 5 
Eueretagrotis attenta (Grt.) 8-29 July 60 
Heptagrotis phyllophora (Grt.) 27 June—25 July 39 
Cryptocala acadiensis (Bethune) 22-24 July 4 
Noctua pronuba L. 23-25 July 3 


1 Identification uncertain, may include or consist entirely of Hypagyrtis unipunctata (Haworth) (Geometridae). 
2 Includes Hydriomena divisaria (Walker) (Geometridae). 

3 Includes Syngrapha abstrusa Eichlin & Cunningham (Noctuidae). 

4 Identification uncertain, may include or consist entirely of Xestia adela Franclemont (Noctuidae). 


Journal of the Lepidopterists’ Society 
48(2), 1994, 106-110 


A REVIEW OF THE PHLOXIPHAGA GROUP OF THE 
GENUS HELIOTHIS (NOCTUIDAE: HELIOTHENTINAE?*) 
WITH DESCRIPTION OF A NEW SPECIES 


DAVID F. HARDWICK 


Research Associate, Centre for Land and Biological Resources Research, 
Agriculture Canada, Ottawa K1A 0C6, Canada 


ABSTRACT. The name Heliothis phloxiphaga at present embraces three distinct 
species: H. phloxiphaga G. & R., which is distributed throughout temperate North Amer- 
ica, H. acesias F. & R., which occurs from Nevada and Idaho eastward to Ontario, and 
H. australis, new species, which is distributed from southern Mexico northward to New 
Mexico and Arizona. 


Additional key words: taxonomy, cryptic species, Heliothis acesias, Heliothis aus- 
tralis, genitalia. 


For several decades, Heliothis acesias F. & R. has been considered 
only a color form of Heliothis phloxiphaga. Although the valvae of the 
male genitalia show no evident differences, the everted vesica of the 
penis of the male and the bursa copulatrix of the female do show 
consistent differences. A third member of the group, a predominantly 
Mexican species, is larger than either H. phloxiphaga or H. acesias and 
also differs from these in the conformation of the basal portion of the 
vesica and of the bursa copulatrix. 


Heliothis phloxiphaga G. & R. 
Figs. 1, 4, 5, 8 


Heliothis phloxiphaga Grote & Robinson (1867:187). 
Heliothis interjacens Grote (1880:30). 


Heliothis phloxiphaga is one of North America’s most common he- 
liothentines; it is distributed throughout the United States and southern 
Canada and feeds on the flower and seeds of a wide variety of plants. 
The species is in flight throughout the summer months. 

The forewing is light brown without a strong yellow suffusion. The 
median shade of the forewing (Fig. 1) angles outward from the trailing 
margin to the reniform spot, then is abruptly angled inward before 
continuing to the costal margin. The central area of the hindwing of 


* Steyskal (1971) clearly demonstrated that Heliothentinae is the correctly derived name for the group. According to 
Opinion 1812 of the International Commission on Zoological Nomenclature, however, the name Heliothinae is to be 
made a conservandum as a subfamily name. This Opinion is without validity. When the submission was made by I. B. 
Nye in 1980, with a supporting addendum by the then-Secretary, R. V. Melville, I submitted a defense of the term 
Heliothentinae. My comments were suppressed by the Secretary and never submitted to the Commissioners. In view of 
this cavalier action, I requested of his successor as Secretary, P. K. Tubbs, that the matter be reopened. He refused. In 
his supporting addendum to Nye’s submission, Melville noted that a noctuid moth, being nocturnal, “would be most 
unlikely to be exposed to the sun,” and that Ochsenheimer’s name was due to a printing error. This, of course, is 
completely fallacious. Obviously, Ochsenheimer was quite aware that the components of his proposed genus were often 
diurnally active. 


VOLUME 48, NUMBER 2 107 


ci, 


Fics. 1-5. Adults and genitalia of Heliothis spp. 1, Heliothis phloxiphaga G. & R., 
Aweme, Manitoba; 2, H. acesias F. & R., Spanish Fork, Utah; 3, H. australis n. sp., 
paratype, Lomas de Chapultepec, D.F., Mexico; 4, Female genitalia of H. phloxiphaga; 
5, Valves of male genitalia of H. phloxiphaga. 


the species is white. In the male genitalia (Figs. 5, 8), there are two 
elongate basal diverticula on the vesica, the one on the right noticeably 
stouter than the one on the left. In the female genitalia, the appendix 
bursae is slightly shorter than the fundus bursae (Fig. 4), and terminates 
in a short, slender, curving tail. 

Expanse (Mean + SD): 34.38 + 1.57 mm (n = 80). 


108 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Heliothis acesias F. & R. 
Figs. 2, 6, 9 


Heliothis acesias Felder & Rogenhofer (1872:pl. 108, fig. 42). 
Heliothis luteitinctus Grote (1875:426). 


The known distribution of Heliothis acesias extends from north- 
eastern Nevada and southern Idaho, northward to southern Alberta, 
thence eastward to southern and eastern Ontario. It has been collected 
on dates between the middle of June and the end of September. 

The species is of comparable size to H. phloxiphaga, but has yellow- 
ish-fawn forewings that are usually somewhat broader for their length 
than those of H. phloxiphaga. The median shade of the forewing (Fig. 
2) is broadly excurved between the trailing and costal margins of the 
wing. The emarginating dots around the reniform and orbicular spots 
are usually more prominent than those of H. phloxiphaga. The central 
area of the hindwing is usually light yellow, less commonly white. In 
the male genitalia (Fig. 9), the base of the vesica is provided with only 
two shallow eversions. In the female, the appendix bursae (Fig. 6) is 
long and recurves anteriorly around the distal end of the fundus bursae. 

Expanse: 33.0 + 1.88 mm (n = 24). 


Heliothis australis Hardwick, new species 


Figs. 3, 7, 10 


Species usually larger than either H. phloxiphaga or H. acesias. Male 
with vestiture of head and thorax varying from reddish brown to olive- 
brown. Forewing light reddish brown, paler centrally than peripherally, 
with a very large and prominent reniform spot (Fig. 3). Median shade 
rather acutely excurved but not usually conspicuous. Central area of 
hind wing medium to dark yellow. In male genitalia, base of vesica 
(Fig. 10) provided with two stout diverticula, the left noticeably longer 
than right. 

Female with maculation and coloring as in the male. Female genitalia 
(Fig. 7) with appendix bursae approximating fundus bursae in length. 

Expanse: 37.2 + 2.46 mm (n = 17). 

Type material. Holotype, male: San Cristobal de las Casas, 7200 ft., 
Chiapas, Mexico, 5 May 1969 (J. E. H. Martin). Paratypes, 20 males, 
6 females: Lomas de Chapultepec, D.F., Mexico, 9 May 1989, male; 
10 June 1940, male; 18 June 1935, male; 28 June 1935 (2 males); 16 
August 1976, male (T. Escalante). San Jacinto, D.F., Mexico, 24 July 
1932, female; 19 October 1932, male; 2 October 1935, male. 24 mi. E 
El Salto, 7500 ft., Durango, Mexico, 15 & 17 July 1964, 2 males (J. E. 
H. Martin). Mexico, Mexico, 2 females. Bent, New Mexico, male, fe- 
male. 13 mi. SE Cloudcroft, 7100 ft., New Mexico, 9 September 1975, 


VOLUME 48, NUMBER 2 109 


A 


Fics. 6-10. Genitalia of Heliothis spp. 6, Female of H. acesias; 7, Female of H. 
australis; 8, Apex of aedeagus and base of vesica of H. phloxiphaga; 9, Aedeagus and 
vesica of H. acesias; 10, Aedeagus and vesica of H. australis. 


male, female (Lafontaine & Bowen). Cedar Crest, Sandia Mts., Ber- 
nalillo County, New Mexico, 4 July 1998, male (R. H. Leuschner). 
Prescott, Arizona, female. Alpine, White Mts., Apache County, Arizona, 
24-25 July 1965, 2 males (R. H. Leuschner). Greer, 8300 ft., White 
Mts., Apache County, Arizona, 4 August 1969, 2 males (R. H. Leusch- 
ner). Springerville,. Apache County, Arizona, 18-24 July 1971, male. 
Two males without data. 

Holotype in the Canadian National Collection. Paratypes in the 
C.N.C., the U.S. National Museum, the Los Angeles County Museum, 
and the collection of R. H. Leuschner. 

Heliothis australis is distributed from the state of Chiapas in southern 
Mexico northward to southern New Mexico and Arizona. The species 
has been collected on dates between early May and mid-October. 


ACKNOWLEDGMENTS 


I am grateful to J. Donald Lafontaine and Eugenie Krelina of the Centre for Land 
and Biological Resources Research for assistance in preparing genitalic slides and arranging 
for the photography of specimens. Julian Donahue and Ron Leuschner provided distri- 
butional data on Heliothis australis. 


LITERATURE CITED 


FELDER, R. & A. F. ROGENHOFER. 1865-1875. Reise der Osterreichischen Fregatte 
Novara um die Erde. Wien. 


GroTE, A. R. 1875. On North American Noctuae. Proc. Acad. Nat. Sci. Philadelphia 
27:418-427. 


1880. Descriptions of new Noctuidae. Bull. Brooklyn Entomol. Soc. 3:29-31. 


110 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


GROTE, A. R. & C. T. ROBINSON. 1867. Descriptions of American Lepidoptera—No. 
2. Trans. Amer. Entomol. Soc. 1:171-192. 


STEYSKAL, G. C. 1971. On the grammar of the name Heliothis Ochsenheimer (Noc- 
tuidae). J. Lepid. Soc. 25:264—265. 


Received for publication 17 July 1998; revised and accepted 24 October 1998. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 111-120 


LIFE HISTORY OF PACHLIOPTA RHODIFER 
(PAPILIONIDAE: TROIDINI) 


K. VEENAKUMARI AND PRASHANTH MOHANRAJ 


Central Agricultural Research Institute, P. B. No. 181, Port Blair, 744101, 
Andaman Islands, India 


ABSTRACT. The Andaman clubtail, Pachliopta rhodifer (Butler), is one of three 
papilionids endemic to the Andaman and Nicobar islands, India. This elegant red-bodied 
swallowtail, with its unique red spatulate tail, previously was known only from the imago. 
We detail the life history of this species and discuss implications for butterfly conservation 
in these islands. 


Additional key words: Aristolochiaceae, butterfly conservation, Andaman Islands, 
Nicobar Islands, India. 


The Andaman and Nicobar islands have been identified as one of 
the provinces of the Indomalayan biogeographic realm requiring urgent 
attention to ensure adequate protection of their distinctive biological 
communities (Anonymous 1985, ICBP 1992). These islands are situated 
in the Bay of Bengal, Indian Ocean, between 6-14°N latitude and 92- 
94°F longitude, and consist of 306 islands. They stretch over 700 km 
and occupy a total area of 8249 km? (Anonymous 1986). The Andamans 
are separated from the Nicobars by the Ten Degree Channel, a distance 
of about 100 km. The two island groups have characteristic biotic 
elements, with each island group harboring a set of unique endemics 
(Anonymous 1986, Rao 1986). 

On the basis of the number of endemics found in a region, Collins 
and Morris (1985) rate the Andaman and Nicobar islands sixteenth out 
of a total of 51 critical swallowtail faunas worldwide. In spite of the 
high priority ranking, no attempt has been made to study the life 
histories, food plants, and other requirements of Graphium epaminon- 
das (Oberthir), Papilio mayo Atkinson, or Pachliopta rhodifer (Butler), 
the three endemic papilionids that are thought to be confined to Great 
Andaman (which includes the three neighboring islands of North, Mid- 
dle, and South Andaman). Even Ferrar, who collected Lepidoptera on 
these islands for eight years, confined himself to the study of adults 
(Ferrar 1948). 

Four species of Pachliopta are known to occur in the fects and 
Nicobar islands. Two of these, P. coon (Fabricius) and P. hector (Lin- 
naeus), are protected by Indian law (Schedule I of the Indian Wildlife 
Protection Act, 1972, which came into effect on 2 October 1980). The 
former species is very rare and found only in the Nicobars in India; 
the latter is rare and confined to the Andamans and mainland India. 
Neither of the other two species, P. rhodifer and P. aristolochiae (F.), 
are considered threatened, even though the status of the endemic P. 


2 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


rhodifer is not clearly known (Collins & Morris 1985). Pachliopta rhodi- 
fer is one of 17 species of the genus listed by Collins and Morris (1985) 
as requiring further monitoring and research. 

We present below the results of our study on the life history of P. 
rhodifer, the only troidine swallowtail endemic to the Andaman Islands. 


Previous Collections 


Ferrar (1948) states that there is a “good series” of P. rhodifer at the 
British Museum (Natural History) and at Calcutta. However, this species 
has proved to be elusive to those who have collected recently on these 
islands (Arora & Nandi 1980). Of nine expeditions from the Zoological 
Survey of India, Calcutta, only four succeeded in collecting this species. 
Cumulatively these surveys covered eight months of the year. They 
collected P. rhodifer during four months, i.e., January, March, April, 
and December. They collected a total of 16 males and four females. 
Nakamotu also succeeded in collecting four males and five females in 
the months of October and November 1979 (Tsukada & Nishiyama 
1982). 

Pachliopta rhodifer was known only from various places in South, 
Middle, and North Andamans (Ferrar 1948, Tsukada & Nishiyama 
1982) until Arora and Nandi (1980) collected a male from Car Nicobar 
in December 1972. This male was perhaps a straggler and may have 
been blown over from S. Andaman to C. Nicobar (150 km). The food 
plant is found in the Nicobar Islands (Rao 1986), hence the butterfly 
could survive and breed there. 

It is perhaps the scarcity of P. rhodifer in eel cies around the 
world that has resulted in its relatively high value in the insect trade 
(Smart & Smart 1982). One male specimen reportedly collected from 
North Andaman was listed for sale for $400 U.S. We believe that rarity 
in collections is more a result of the stringent regulations imposed by 
the Government of India rather than a reflection of the rarity of the 
butterfly. 


MATERIALS AND METHODS 


We discovered a last instar larva of P. rhodifer during a collecting 
trip to the forests of South Andaman in late October 1990. The discovery 
of the host of P. rhodifer was the key to searching for the early stages. 
Over a two-year period we visually searched mainly three localities for 
early stages: Garacharma, Mt. Harriet, and Chiciy ata all on the 
island of S. Andaman. 

Upon locating host plants, which have a tendency to grow in patches, 
a thorough examination for eggs, larvae, and pupae was conducted. All 


VOLUME 48, NUMBER 2 1138 


early stages discovered in the field were brought into the laboratory 
and housed in transparent, plastic jars of variable dimensions—the first 
and second instars in the smallest (6.5 cm high x 5 cm diameter), the 
third to fifth instars in medium (10 cm high x 7 cm diameter), and 
the final instar in the largest (19 cm high x 11 cm diameter) containers. 
Food (detached, whole, tender leaves) was supplied fresh every day. 
The rearing containers also were cleaned daily of all fecal matter and 
old food. Moisture that settled on the walls of the containers was wiped 
away with a dry cloth. A dry twig, slightly less than the height of the 
container, was placed diagonally in containers with final instars to 
facilitate pupation. All rearings were carried out in the laboratory at 
ambient temperature (23-30°C) and humidity (79%). Adults were re- 
leased back into their habitat. 

The study was conducted primarily on the main campus of the 
Central Agricultural Research Institute at Garacharma, approximately 
6 km southwest of Port Blair on the island of S. Andaman, India. Patches 
of forest that have been retained in parts of the campus were found to 
support populations of the butterfly. Although subject to intense human 
intrusion, these patches continue to retain a number of plant and animal 
species that were a part of the original forest that once covered this 
area (Fig. 1). The larval food plant of P. rhodifer was found in a number 
of patches, some of which suffered more human damage than others. 
A large population of the immature stages was present in a relatively 
disturbed area where the larval food plant grew in the shrub layer close 
to the ground beneath the canopies of coconut, oil palm, and other 
trees. This area was subject to periodic weeding operations. 

Additional observations were made during periodic visits to Chidi- 
yatapu and Mt. Harriet, both situated in the southern half of S. An- 
daman. Although subject to less anthropogenic activities than the Gar- 
acharma site, these sites yielded fewer eggs and larvae of P. rhodifer, 
despite the fact that patches of the larval food plant at both these sites 
were more dense and healthier in appearance. 


RESULTS 


Thottea tomentosa (Blume) Ding Hou (Aristolochiaceae), a low, 
woody, creeping under-shrub in the forests, was found to be the sole 
food plant of P. rhodifer (Figs. 1 & 2). The maximum height of the 
plant was 51.0 cm. Only the younger, densely tomentose terminal leaves 
were eaten; the lower older leaves were never fed upon. The older 
leaves were tougher and had a lower percentage of water than the 
younger leaves. After consuming most or all of the young, tender leaves 
of a plant, the larvae leave to find a new host plant. 

We observed adults feeding on the flowers of Vitex trifolia L. (Ver- 


114 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


benaceae), Acacia sp. (Mimosaceae), Ixora sp. (Rubiaceae) and Intsia 
bijuga (Coleb.) O.K. (Caesalpineaceae). 


Egg (Fig. 3): Similar to other troidine eggs; pale orange with a smooth circular area 
on top. 

First instar: Pale orange, partly suffused with black. Head, prothoracic shield, and anal 
legs deep glossy black; osmeterium pale orange. Tops of tubercles capped with about a 
dozen long, black setae. [This is characteristic of the early instars and unlike that of later 
instars.] Entire surface of body covered with minute hairs. Head capsule black in all 
instars. 

Second instar: All larvae with similar color pattern from this instar on. The only 
difference between this and the following instars is that the red bands are incomplete 
along the mid-dorsal line, resulting in a continuous black mid-dorsal line, obliterated in 
the latter instars. 

Third through sixth instars (Figs. 4 & 5): Rich purple-black, studded with intense 
orange-red and a few purple-black tubercles, interspersed with narrow bright orange-red 
bands running along the posterior margins of abdominal segments IV and VII. Head 
glossy black with a number of stiff bristlelike black setae. Clypeus dirty white and 
translucent. 

On either side of the mid-dorsal line a row of bright-red tubercles on all segments 
except abdominal segments II, III, V, and VI, which are intense purple-black. A sub- 
dorsal row of orange-red tubercles on thoracic segments only, this row absent on the 
abdominal segments. A subspiracular row of tubercles on all thoracic and abdominal 
segments, orange-red except on abdominal segments V and VI where they are black. A 
complete row of sub-ventral tubercles, all bright orange-red except that on the last 
abdominal segment which is claret. A single small ventral tubercle exclusive to the second 
abdominal segment. All tubercles covered with setae, black on the red tubercles, silver- 
grey on the black tubercles. Ventral surface black, suffused with claret. 

Spiracles black, outlined in white. Spiracles on abdominal segment IV in the black area 
apposite, but posterior to, the orange-red band. Spiracles on abdominal segments I and 
II progressively dorsal than those on all other abdominal segments, which form a straight 
line. Consequently, the subspiracular tubercles also have the I abdominal one highest, 
followed a little lower by the II abdominal tubercle; the remaining tubercles occur in a 
similar position on each segment, forming a straight line. Distances between the subspi- 
racular and sub-ventral tubercles are variable in the anterior segments; the prothoracic 
and all abdominal tubercles of the two rows very close, their bases almost touching; the 
two spiracles on the meso- and metathoracic segments relatively further apart. The only 
tubercles on the prothorax are the subspiracular and the sub-ventral, which are deep 
orange-red. Dorsal space between the subspiracular tubercles covered by a light, orange- 
red smooth pad, faintly grooved along its mid-dorsal line. Black prothoracic shield behind 
this pad. A short red lateral band posteriorly, on the IIJ abdominal segment starting from 
the base of the subspiracular tubercle and extending up to about % the distance to the 
dorsal tubercle. Red band on the IV abdominal segment posteriorly notched on the mid- 
dorsal line. Red band on abdominal segment VII divided mid-dorsally by a rich purple- 
black band. Depressed black spot in red bands of abdominal segments IV (above the base 
of the subspiracular tubercle) and VII (antero-dorsally at base of the subspiracular tu- 
bercle). 

Pupa (Fig. 6): Orange-brown, resembling a dry leaf. Girdle black; cream lateral mark- 
ings anterior to girdle. Two red, glistening, triangular areas on either side of the mid- 
dorsal line on anterior end of dorsal area of thorax. Blunt, dorsal horn with faint dorsal 
red line and foliaceous extensions/carinae from antero-ventral region, extending laterally 
to abdominal segment I. Circular, red spot in mid-dorsal region of abdominal segment 
II. Two orange-red areas or irregular spots on abdominal segment III laterally. Sparse, 
minute hairs in pits on pupal surface, increasing in number around dorsal end of spiracles. 
Additional pair of processes on either side of dorsal horn and on lateral margins of 
abdominal segment I. 


VOLUME 48, NUMBER 2 115 


Fics. 1-7. Habitat and early stages of Pachliopta rhodifer. 1, Habitat of P. rhodifer; 
2, Thottea tomentosa, the larval food plant, growing at the base of a tree; 3, Egg of P. 
rhodifer, 4, Third instar of P. rhodifer; 5, Fifth instar of P. rhodifer; 6, Pupa of P. 
rhodifer; 7, Adult male of P. rhodifer. 


Eggs are laid singly. Females fly low over the undershrub layer, 
alighting on vegetation frequently, including the host plant, but do not 
often lay eggs. When laying an egg, the female alights on the host 
plant, bends her abdomen beneath the leaf surface, and lays an egg 
along or on one of the veins of a young leaf, but usually not on the 
midrib. Eggs generally are laid on the under surface of leaves of the 


116 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


TABLE 1. Duration and size of the pre-imaginal stages of P. rhodifer. 


Eggs I Il Ill IV V VI PP P 


Duration (days) 


n 2 4 7 13 16 18 19 19 13 
Range >6 2-4 2-4 3-6 4-8 o-7 7-14 1-4 15-64 
Mean — 3 2.83 3.62 5.138 One 9.63 1.95 26.69 
SE — 0.41 0.26 0.25 0.24 0.16 0.39 0.14 5.19 
Size (cm) 

n — 2 6 13 16 18 20 — 19 
Range — 0.7-0.9 0.6-1.0 0.6-1.5 1.8-2.2 2.3-3.1 2.8-3.7 — 3.1-3.8 
Mean — 0.8 0.85 1.16 1.67 2.68 3.31 — 3.44 
SE — 0.1 0.06 0.07 0.06 0.06 0.07 — 0.04 


* Total life cycle: Mean from I instar to adult = 58.6 days = 59 days. Mean from egg to adult = over 64.6 days = 
over 65 days. 


host plant or rarely on adjacent vegetation. Because the undersurfaces 
of Thottea leaves generally are encrusted with a layer of mud, the 
veins are the only places where eggs can be laid safely without being 
dislodged. Only one egg is laid on each leaf. On one occasion, one egg 
was found on a blade of grass; another was found on a cane leaf. The 
incubation period is at least six days, after which the first instar larvae 
emerge. Larvae eat the chorion and begin to feed on the tender young 
leaves (see Table 1 for sizes and durations of all stages). 

The first instar larva appears to remain on the leaf on which it hatches; 
later instars move not only from one leaf to another on the same plant, 
but also from plant to plant. Later instars have been observed moving 
along the ground, probably in search of a new host plant; host plants 
tend to have clumped distributions. Larvae are forced to move from 
plant to plant because they feed exclusively on the new flush. 

Although we were able to find all stages from eggs to last larval instar, 
we never discovered prepupae or pupae, suggesting that pupation oc- 
curs away from the host plant. 

The prepupal period usually lasts about two days but in one case it 
lasted four days. 

The total life cycle lasts about 65 days. As the eggs were collected 
from the field, and not from females bred in the laboratory, it was not 
possible to ascertain when they had been laid, leading to an uncertainty 


TABLE 2. Duration and head capsule measurements of P. rhodifer larva when passing 
through seven instars (n = 1). 


Duration (days) >8 8 4 4 5 5, 4 lea 10 
Width of head capsule (mm) — 0.9 1D ef DS 2.9 3.9 — 


VOLUME 48, NUMBER 2 : | BIE, 


TABLE 3. Mortality pattern in captive pre-imaginal stages of P. rhodifer in S. An- 
daman. 


Ege I Il TH IV V VI PP P 
Total collected 5) 4 iT 13 16 19 I 23 23 
No. dead 3 = == a a 1 ] 3 9 


Percent mortality 60 


in the incubation period and consequently in the number of days taken 
to complete the entire life cycle. 

When we searched for the eggs and larvae of P. rhodifer in April, 
the driest month in the year, we could find only one egg of the butterfly 
and no new flush of the food plant. On hatching, the larva had to be 
fed older, poor quality leaves. This individual passed through seven 
larval stages. Nevertheless, the total larval period did not vary from 
those rearings that had six instars. Dimensions of the head capsules and 
the durations of larval instars of this individual are presented in Table 
2. 

Two eggs and a third instar larva collected in the field were para- 
sitized. The egg parasitoid was identified as Telenomus (Aholcus) sp. 
(Scelionidae: Hymenoptera). The larval parasitoid could not be iden- 
tified, as the hymenopteran adults failed to emerge from their cocoons. 
A certain degree of mortality (Table 3) also was noticed while rearing 
the butterfly. From the first to fourth instar, all larvae survived. Mor- 
tality occurred in the egg stage and in larval instars V and VI, as well 
as during the prepupal and pupal stages. Mortality was highest in the 
egg and pupal stages. 

One instance of mating in flight was observed at about 1100 h in the 
Garacharma farm of the C.A.R.I. campus when a pair in copula flew 
at a height of about 5-6 meters and alighted on the branch of a tree 
well beyond our reach. A mating pair also was seen at about 1700 h 
on a blade of grass along an earthen embankment at the base of Mt. 
Harriet. 


DISCUSSION 


Thirteen genera in the tribe Troidini feed on aristolochiaceous food 
plants during their larval stages (Igarashi 1984). Of the four species of 
Pachliopta found in the Andaman and Nicobar islands, two are rare 
and their local food plants remain unknown. The other two species, P. 
aristolochiae and P. rhodifer (Fig. 7), feed exclusively on Thottea 
tomentosa. Thottea tomentosa, though only locally abundant, is much 
more frequent in its occurrence than Aristolochia tagala Cham., the 
only other member of the Aristolochiaceae found on these islands (Ding 


118 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Hou 1981, Rao 1986). Although we found P. rhodifer and P. aristo- 
lochiae occurring together in our study sites, we did not find them in 
numbers large enough to suggest intra-specific competition during the 
larval stage. Therefore we feel that there are no specific larval adap- 
tations that have arisen as a result of competitive interactions. It is 
interesting to note that although Thottea is widely distributed in the 
Malesian region and on the Indian Subcontinent (Ding Hou 1981), P. 
rhodifer has failed to expand its range. In fact, although T. tomentosa 
occurs in both the Andaman and Nicobar islands (Rao 1986, specimens 
at the Botanical Survey of India herbarium, Port Blair), P. rhodifer 
apparently is restricted to Great Andaman [with the exception of the 
single stray record of the species from Car Nicobar by Arora and Nandi 
(1980)]. 

Open water between islands may be an effective barrier preventing 
dispersal of this species. Though Tsukada and Nishiyama (1982) indicate 
that it is an exceedingly slow flyer, our observations reveal that the 
species is capable of fairly fast (though not vigorous), bobbing flight 
when disturbed. 

The presence of six larval instars, instead of the usual five as in all 
other Papilionidae, lends credence to classifying P. rhodifer as a distinct 
species and not as a form of P. coon (Miller 1987). Although the larvae 
may pass through an additional instar when under stress (possibly be- 
cause of food of inferior quality), six instars is the norm for P. rhodifer. 

Like all troidines, P. rhodifer has red-tuberculate, Aristolochia-feed- 
ing larvae that are aposematically colored and probably unpalatable to 
predators (Hancock 1983, 1988, DeVries 1987). Because we found lar- 
vae during most months of the year, we suspect that it is multivoltine, 
as in other Pachliopta species (Igarashi 1984). Pachliopta rhodifer lays 
eggs singly and the larvae are solitary feeders, thus conforming to the 
pattern in other Pachliopta (Igarashi 1984). 

Tsukada and Nishiyama (1982) state that adults of P. rhodifer ‘be- 
come active only at dusk and fly toward 5 p. m.’ Our studies failed to 
corroborate this observation. We observed P. rhodifer in flight in the 
morning and in the evening (from 0700 h to 1800 h). 

In the Oriental region, only the peripheral areas such as the Andaman 
Islands are rich in endemic Papilionidae (Hancock 1983). Williams et 
al (1991) suggest the incorporation of the extent of geneological dif- 
ferences into the biodiversity measure in addition to the endemicity 
criterion. If this is done, the Andaman Islands are likely to move further 
up the priority list presented by Collins and Morris (1985) because the 
three endemic papilionids in the Andamans belong to three different 
tribes—Leptocircini, Papilionini, Troidini (Hancock 1983). 

This diversity of endemics, together with the fact that all endemics 


VOLUME 48, NUMBER 2 119 


so far have been reported almost exclusively from Great Andaman, 
indicates that butterfly conservation in the region is extremely impor- 
tant. The Great Andaman, which makes up well over half the area of 
the Andaman Islands (constituting about 70 percent of the total land 
area of the Andamans) has been facing great pressure from human 
activities. Since the mid-1800’s it has been increasingly cleared of its 
natural vegetation to meet the growing demands of a rapidly expanding 
human population that favors settlements close to Port Blair, the capital 
of these islands. The food plants of at least two endemic species of 
Papilionidae, Graphium epaminondas and P. rhodifer, both of which 
have patchy distributions in the forests of these islands, may be facing 
a threat from these activities of man. There is thus an urgent need for 
studies of the type being conducted by Pollard (1977) to assess the status 
of the endemic species of Lepidoptera, particularly when the clamor 
for the ‘development’ of these islands through increased urbanization 
and industrialization is on the rise. 


ACKNOWLEDGMENTS 


We are extremely grateful to A. K. Bandyopadhyay, Director, Central Agril. Res. Inst., 
Port Blair, for having been a constant source of encouragement to us in an environment 
in which marked distinctions are made between pure and applied research. 

We thank E. Tata Rao, S. Sree Latha, and Omana and Mukesh Ram for help in the 
field. We are grateful to B. Pardhasardhi for the photographs. We owe special thanks to 
N. M. Collins, J. D. Holloway, R. I. Vane-Wright, and H. Smith for their interest in our 
work and for their comments on the manuscript which greatly helped improve it. We 
thank A. Polaszek for identifying the egg parasitoid and M. K. Vasudeva Rao for iden- 
tifying the host plant. We also appreciate the care taken by the two anonymous referees 
in going through the original manuscript which has resulted in this improved final version. 
We also are thankful to T. V. R. S. Sharma, M. Prem Kumar and Janet Camp for help 
in the preparation of the manuscript. 


LITERATURE CITED 


ANONYMOUS. 1985. The Corbett Action Plan for protected areas of the Indomalayan 
realm. IUCN Commission on National Parks and Protected Areas. Corbett National 
Park, India. 

1986. An integrated environmentally sound development strategy for the An- 
daman and Nicobar islands. GOI, N. Delhi. 

AroRA, G. S. & D. N. NANDI. 1980. On the butterfly fauna of Andaman and Nicobar 
islands (India). I. Papilionidae. Rec. Zool. Surv. India 77:141-151. 

COLLINS, N. M. & M. G. Morris. 1985. Threatened swailowtail butterflies of the world. 
The IUCN Red Data Book. IUCN. Gland, Switzerland. 401 pp. 

DEVBRIES, P. J. 1987. The butterflies of Costa Rica and their natural history: Papilionidae, 
Pieridae, Nymphalidae. Princeton Univ. Press, New Jersey. 327 pp. 

DING Hou. 1981. Florae Malesianae praecursores LXII on the genus Thottea (Aristo- 
lochiaceae). Blumea 27:301-332. 

FERRAR, M. L. 1948. The butterflies of the Andamans and Nicobars. J. Bombay Nat. 
Hist. Soc. 47:470-491. 5 pls. 

HANCOCK, D. L. 1988. Classification of the Papilionidae (Lepidoptera): A phylogenetic 
approach. Smithersia 2:1—48. 


120 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


1988. A revised classification of the genus Atrophaneura Reakirt (Lepidoptera: 
Papilionidae). Aust. Ent. Mag. 15(1):7-16. 

ICBP. 1992. Putting biodiversity on the map: Priority areas for global conservation. 
ICBP, Cambridge. 

IGARASHI, S. 1984. The classification of the Papilionidae mainly based on the morphology 
of their immature stages. Tyo to Ga 34(2):41-96. 

MILLER, J.S. 1987. Phylogenetic studies in the Papilionidae (Lepidoptera: Papilionidae). 
Bull. Amer. Mus. Nat. Hist. 186:365-512. 

POLLARD, E. 1977. A method for assessing changes in abundance of butterflies. Biol. 
Conserv. 12:115-134. 

Rao, M. K. V. 1986. A preliminary report on the angiosperms of Andaman & Nicobar 
islands. J. Econ. Tax. Bot. 8(1):107-184. 

SMART, P. & G. SMART. 1982. Catalogue. St. Mary’s, Bramber, Nr. Steyning, W. Sussex, 
England. 

TSUKADA, E. & Y. NISHIYAMA. 1982. Butterflies of the Southeast Asian islands. Vol. I. 
Papilionidae (transl. K. Morishita). Plapac Co. Ltd., Tokyo. 457 pp. 

WILLIAMS, P. H., C. J. HUMPHRIES & R. I. VANE-WRIGHT. 1991. Measuring biodiversity: 
Taxonomic relatedness for conservation priorities. Aust. Syst. Bot. 4:665—-679. 


Received for publication 25 January 1993; revised and accepted 2 October 1993. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 121-127 


RECENT INTRODUCTIONS OF RIPARIAN NOCTUID 
MOTHS FROM THE PALAEARCTIC REGION TO 
NORTH AMERICA, WITH THE FIRST REPORT OF 
APAMEA UNANIMIS (HUBNER) 
(NOCTUIDAE: AMPHIPYRINAE) 


KAURI MIKKOLA 


Finnish Museum of Natural History, P.O. Box 17, 
FIN-00014 University of Helsinki, Finland 


AND 


J. DONALD LAFONTAINE 


Centre for Land and Biological Resources Research, 
Ottawa, Ontario K1A 0C6, Canada 


ABSTRACT. The Palaearctic moth Apamea unanimis (Hiibner) (Noctuidae: Am- 
phipyrinae) is reported as new to North America on the basis of a female specimen from 
the vicinity of Ottawa, Ontario, Canada, collected 11 June 1991. The larva of this species 
feeds on riparian grasses such as Phragmites and Phalaris (Poaceae). Two other recent 
introductions of Noctuidae from the Palaearctic to North America, Apamea ophiogramma 
(Esper) and Rhizedra lutosa (Hiibner) (Amphipyrinae), both apparently established in 
several areas, feed as larvae on these same plants. The apparent synchronous establishment 
of three Palaearctic species with similar ecological associations suggests that they may 
have arrived by similar means, possibly aided by the modern shipping practices. 


Additional key words: Rhizedra lutosa, Apamea ophiogramma, Phragmites, Phal- 
aris. 


Among the 95 species of Noctuidae recently listed as Holarctic by 
Mikkola et al. (1991), 26 were considered as probable introductions by 
man or assisted by man. Thirteen are common European species, prob- 
ably transported from Europe to the west (or over the Pacific). No 
corresponding introductions from North America to Europe are known. 
Possible reasons for this bias are introduced European plants providing 
host plants in North America, but not vice versa, and the possible 
saturation of European agroecosystems (Mikkola et al. 1991). 

The two most recent introductions differ from earlier ones in that 
the species are associated with moist habitats (Mikkola et al. 1991). 
Rhizedra lutosa (Hiibner) (Figs. 6-8) was reported by McCabe and 
Schweitzer (1991) from New Jersey, and Apamea ophiogramma (Esper) 
(Figs. 4,5) by Troubridge et al. (1992) from the Vancouver area, British 
Columbia. We list several new localities for both of these species from 
eastern North America. 

We would like to draw the attention of North American lepidopterists 
to these two species and, in particular, to a third amphipyrine species 
first found in North America in 1991. 


122 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Apamea unanimis (Htbner, 1813) 


The first North American specimen was collected in Pinhey Forest 
Preserve, Nepean, 12 km SW of Ottawa on 11 June, 1991 (J. D. La- 
fontaine). This female specimen (Fig. 1) was caught at light in a garden 
at the margin of a mixed forest, in the garden there is a small artificial 
pond with tall grasses and sedges transported from nearby ditches. 

Apamea unanimis is a small Apamea with a wingspan (i.e., distance 
of forewing outer margins in a well spread specimen) of 29 to 38 mm 
(mean 30.9 mméé, 31.7 mm; the difference is significant, Mikkola & 
Jalas 1979). It is a highly variable species, and the forewing coloration 
varies from pale luteous gray brown with a darker costal area to uni- 
colorous blackish brown (cf. Figs. 1-8). The median area is weakly 
marked. The presence of fine white lining on the outer side of the 
reniform spot is a good specific character. 

The species is not closely related to any North American Apamea 
species but is most likely to be misidentified as a dark form of Apamea 
remissa indocilis (Walker). It can be distinguished from remissa by its 
white-lined reniform spot. The most obvious feature distinctive from 
“Oligia”’ fractilinea (Grote), and from the Palaearctic Mesapamea 
secalis (Linnaeus), is the large hindwing discal spot (well visible on the 
underside of the wing). 

The determination of male specimens of A. unanimis may be easily 
confirmed by brushing the valva. The cucullus is upright as in most 
Apameas, and a large sclerotized, coiled digitus is present at the anterior 
lower corner. 

Apamea unanimis occurs from the Pyrenees and northern Italy to 
the central parts of Fennoscandia (about the 63rd parallel) in Europe, 
and from there the distribution extends through Russia, Caucasus, and 
Siberia to the Amur River. The species is mainly rare and localized. 
However, in suitable habitats, such as rocky sea-shores, the species may 
be abundant in June and early July. It can be collected most easily by 
sugaring. 

The larva feeds between the leaves of riparian grasses such as Phrag- 
mites, Phalaris, and Glyceria overwintering fully grown and pupating 
in the spring inside broken stems (Mikkola & Jalas 1979). The larva 
overwinters in grass tussocks, under loose bark, and in rotten wood, and 
pupates in the early spring in a compact cocoon in the soil (Bretherton 
et al. 1983). 


Apamea ophiogramma (Esper, 1793) 


Apamea ophiogramma was first reported from Langley, British Co- 
lumbia; 4 specimens were collected at light in 1989, 19 in 1990 and 38 
in 1991, between late June and late August, all of them of the typical 


VOLUME 48, NUMBER 2 123 


Fics. 1-8. Specimens of Apamea unanimis (Hubner), A. ophiogramma (Esper) and 
Rhizedra lutosa (Hiibner) from North America and Europe. 1-3, A. unanimis—1, 
female, Canada, Ontario, Ottawa 12 km SW, Pinhey Forest, 11 June 1991, leg. J. D. 
Lafontaine; 2, male, Finland, U: Porvoo, 8 June 1930, leg. E. Suomalainen; 3, female, 
Finland, U: Hanko, 18 June 1983, leg. Nupponen; 4-5, A. ophiogramma—4, female, 
Canada, B.C., 5 km E Langley, 24-30 June 1990, leg. J. Troubridge; 5, male form 
“maerens’, U: Tvérminne, 9 August 1981, leg. A. Jarvela; 6-8, R. lutosa—6, male, 
Finland, U: Espoo, 16 September 1960, leg. P. Utrio; 7, female, U.S.A., N.J., Cumberland 
Co., Port Norris, salt marsh, 30 September 1988, leg. D. Schweitzer; 8, female, England, 
6 September 1892, leg. T. Tunstall. 


124 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


(pale) form (Troubridge et al. 1992). We know of the following spec- 
imens from eastern North America: 


—Vermont, South Burlington, 28 July 1991, J. R. Graham, @ f. ““mae- 
rens”’ 

—New York, Albany Co., Pinebush, 1991, T. L. McCabe, 1 ex. f. 
“maerens”’ 

—Quebec, St.-Jean de Martha, 26 July 1992, T. Thouin, é f. “typica” 

—New Brunswick, Edmundston, 7 August 1992, H. Hensel, 4 f. “mae- 
rens’’. 


Apamea ophiogramma is a small noctuid with a wingspan of 26 to 
32.5 mm. It is easily recognized by the snake-like line between the 
blackish anterior and the pale brownish posterior part of the forewing. 
The forewing groundcolor is smoky gray in the melanic form “mae- 
rens”’ Staudinger, and the contrast of the “‘snake line” is correspondingly 
weaker (Fig. 5). The melanic form in Helsinki, Finland constitutes about 
40 percent of the population (Mikkola & Jalas 1979), but this form is 
not mentioned from Japan (Sugi 1982). The male genitalia of the species 
were illustrated by Troubridge et al. (1992). Sugi (1982) moved the 
taxon to the genus Oligia, but we prefer to retain it in Apamea until 
a generic revision of the Apameini is completed. 

The species is distributed from western Europe to Japan. The larva 
feeds on shore grasses, such as Phragmites, Phalaris and Glyceria (e.g., 
Mikkola & Jalas 1979), living within the stems, leaving them to over- 
winter in the soil, and returning to the stems in the spring. It pupates 
in the soil (Bretherton et al. 1983). The moths fly mainly from mid- 
July to mid-August, and they can be collected near shores and wetlands 
at light or by sugaring. 


Rhizedra lutosa (Htibner, 1803) 


This species was first reported in North America from the salt marshes 
of Delaware Bay, Cumberland County, New Jersey (McCabe & 
Schweitzer 1991) where 18 moths (584 1392) were collected in 1988 and 
1989. The extreme dates were 30 September and 4 November. We 
now report two additional collections: 


—New York, Albany Co., Pinebush, 1991, T. L. McCabe 
—New York, Catskill, West Shokan, 1991, J. G. Franclemont. 


This is a bulky fall-flying species that can be recognized easily from 
the straw-colored, apically sharp forewing. The groundcolor may have 
a reddish-brown hue. The wingspan varies in Finland from 34 to 52.5 
mm, the females being on average larger than the males, and the 
forewing is “almost always characteristically irrorate with black scales” 


VOLUME 48, NUMBER 2 125 


(Mikkola & Jalas 1979). The specimen from New Jersey (Fig. 7) is 
uncharacteristically large and pale. The handbooks from Central Eu- 
rope illustrate similar unicolorously pale moths (e.g., Bretherton et al. 
1988). The male genitalia were illustrated by McCabe and Schweitzer 
(1991). 

The species is distributed from western Europe to Tibet and Japan. 
In Europe, R. lutosa is thought to be monophagous on Phragmites 
australis. The larva lives inside the stem bases and rhizomes of reeds 
that grow on dry land, causing blanching of the leaves; it pupates in 
the reed-humus amongst the rhizomes; the egg overwinters (Mikkola 
& Jalas 1977, Bretherton et al. 1983). The moths fly mainly from late 
August to early October, in and near reeds but, as in many other noctuids 
of this habitat, the females may disperse far from suitable habitat. The 
moths are attracted to light but rarely come to sugar baits. 


DISCUSSION 


The Canadian record of Apamea unanimis fits well with the Eu- 
ropean flight time because in Europe the species is one of the earliest 
Apameas to appear in late spring and early summer. The around 5 
yards long garden pond hardly supports a permanent population of A. 
unanimis, but the species could be established along the shores of 
Ottawa and Rideau rivers, the latter two miles away, where Phragmites 
grows. The other two introduced riparian species seem to be relatively 
widespread, but the sudden accumulation of new observations suggests 
that they are newcomers. 

The external appearance of the moths, combined with geographical 
factors, permits some inferences about the possible sources of the found- 
er individuals. The monomorphic population of Apamea ophiogramma 
in western North America evidently has been imported from the Far- 
East or Japan where the melanic form seems not to occur (there occur 
wild type females in eastern North America, too, but importation from 
there seems improbable). As expected, the eastern population might 
have originated in the industrialized areas of Western Europe (the 
founder could have been a single heterozygous female which would 
have produced both pale and melanic forms). The very pale appearance 
of the American specimens of Rhizedra lutosa points to an origin in 
Great Britain or in continental Central Europe. 

It is curious that three species associated with riparian grasses should 
appear in North America in such a short period. It seems possible that 
modern shipping practices may have contributed to the spread of these 
species. Because one of the species may overwinter as a pupa within 
Phragmites stems or close to them, the other probably lays overwin- 
tering eggs on Phragmites, and the third spends the winter as a medium- 


126 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


7 


ny 


Fic. 9. Distribution of collecting localities of the three European Apameini moths in 
North America. % = Apamea unanimis; @ = Apamea ophiogramma, half-filled = typical 
form, filled = melanic form; @ = Rhizedra lutosa. 


sized larva associated with riparian grasses, it is possible that all three 
species have been transported with Phragmites, or with other riparian 
vegetation. 

Soils are no longer used as ship ballast, but new shipping techniques 
may provide clues. Containers are kept in large storage yards in shore 
areas, and when they are transported into the ships, grass turfs may 
enter the ships. On the other hand, the reproductive strategy of Phrag- 
mites may enhance introduction. The plant mainly reproduces asex- 
ually from the rhizomes, and these often form dense mats along the 
shore. In the winter, the ice movements remove such mats, and rafts 
several meters in length can be seen on or near the shores. Pieces of 
them may enter ships through the open gates. Thus, container ships 
may transport reed material across the ocean, both from Europe to 
eastern North America, and from eastern Asia to the west coast of North 
America. 

If the appearance of these three riparian species reflects a similar 
history of introduction, other Palaearctic riparian species could be ex- 
pected to appear in North America, including the following species 


VOLUME 48, NUMBER 2 27 


feeding on Phragmites as larvae: Archanara spp., Chilodes maritima 
(Tauscher), Arenostola phragmitidis (Hubner), Photedes brevilinea 
(Fenn), Mythimna pudorina (Denis & Schiffermiller), M. straminea 
(Treitschke), M. obsoleta (Hiibner) and Senta flammea (Curtis). Species 
of other lepidopteran families living as larvae on Phragmites or on 
other riparian grasses also may be introduced. 


ACKNOWLEDGMENTS 


We thank J. G. Franclemont (Ithaca, New York), J. R. Graham (South Burlington, 
Vermont), H. Hensel (Edmundston, New Brunswick), T. L. McCabe (Albany, New York), 
D. F. Schweitzer (Port Norris, New Jersey), and T. Thouin (St.-Jean de Martha, Quebec) 
for unpublished observations, and B. Landry, A. Pucat, and R. A. Bracken (Ottawa, 
Ontario) for commenting on the manuscript. 


LITERATURE CITED 


BRETHERTON, R. F., B. GOATER & R. I. LORIMER. 1983. Noctuidae: Cuculliinae to 
Hypeninae, pp. 36-413. In Heath, J. & A. Maitland Emmet (eds.), The moths and 
butterflies of Great Britain and Ireland. Harley Books, Colchester, England. 459 pp. 

McCaBE, T. & D. SCHWEITZER. 1991. Rhizedra lutosa (Lepidoptera: Noctuidae) newly 
introduced to North America. Entomol. News 102:130-132. 

MIKKOLA, K. & I. JALAS. 1979. Suomen perhoset. Y6kk6set 2. (Finnish Lepidoptera. 
Noctuidae 2.). Otava, Helsinki, Finland. 304 pp. 

MIKKOLA, K., J. D. LAFONTAINE & V. S. KONONENKO. 1991. Zoogeography of the 
Holarctic species of the Noctuidae (Lepidoptera): Importance of the Beringian refuge. 
Entomol. Fennica 2:157-178. 

SuGI, S. 1982. Noctuidae. In Inoue, H., S. Sugi, H. Kuroko, S. Moriuti & A. Kawabe 
(eds.), Moths of Japan. Kodansha, Tokyo. 966 + 552 pp. 

TROUBRIDGE, J. T., S. M. FITZPATRICK & J. D. LAFONTAINE. 1992. Apamea ophio- 
gramma (Esper), a palearctic cutworm new to North America (Lepidoptera: Noc- 
tuidae). Canad. Entomol. 124:109-112. 


Received for publication 6 June 1998; revised and accepted 1 October 1993. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 128-147 


SYSTEMATIC POSITION OF SIBATANIOZEPHYRUS AND 
DESCRIPTION OF A NEW SPECIES FROM TAIWAN 
(LYCAENIDAE: THECLINAE) 


YU-FENG Hsu 


Department of Entomological Sciences, University of California, 
Berkeley, California 94720, U.S.A. 


AND 


MING-YAO LIN 
SF, No. 8, Alley 52, Szuwei Road, Taipei, Taiwan, R.O.C. 


ABSTRACT. The systematic position of the Fagus-feeding genus Sibataniozephyrus 
was examined using cladistic methodology. This formerly monotypic genus, confined to 
eastern Asia, previously was lumped with Quercusia, a monotypic genus confined to 
Europe. If the two genera are combined, they form a paraphyletic taxon in the preferred 
cladogram. Hence, this analysis provides support for maintaining the two as distinct 
genera. Sibataniozephyrus kuafui, new species, is described and illustrated from Taiwan. 
The discovery of the new species has interesting biogeographic interpretations. 


Additional key words: Quercusia, Fagus-feeding, Theclini, China, Japan. 


Members of Sibataniozephyrus are moderate sized (forewing length 
15.0-18.6 mm) Theclini “hairstreak’’ butterflies that live exclusively in 
beech (Fagus spp.; Fagaceae) forests. Males spend the majority of their 
adult life flying high around the canopy of beech trees, showing their 
magnificent metallic blue colors. In contrast, females sit motionless most 
of the time, perched in the vegetation, with their somber brown wings 
folded. The larvae of Sibataniozephyrus feed exclusively on soft new 
foliage of the host, which is available only in early spring. As a result 
of the narrow availability of appropriate larval food, members of Si- 
bataniozephyrus are univoltine. 

The taxonomic status of Sibataniozephyris has been the subject of 
considerable controversy. Prior to the discovery of the new species 
described below, the genus was considered monotypic, accommodating 
the single species S. fujisanus (Matsumura), which is restricted to Japan. 
On the basis of similarities in the female genitalia, Sibataniozephyrus 
was lumped by Shirézu and Yamamoto (1956) with the monotypic 
genus Quercusia, which includes the single species Q. quercus (Lin- 
neaus) restricted to Europe. Shir6zu and Yamamoto (1956) did not 
dismiss the possibility that the two may require different genera. Al- 
though the female genitalia of the two are similar, other characters 
(i.e., male genitalia and relative length of the discoidal cell of the 
forewing) are moderately divergent, suggesting that the two genera 
may not form a monophyletic lineage. This taxonomic difficulty has 
been manifested in nearly every treatment that included Sibataniozeph- 


VOLUME 48, NUMBER 2 | 129 


TABLE 1. Number of species of Fagus, Sibataniozephyrus, and Theclini represented 
in the world with special reference to Asia: EU, Europe; AC, mainland China; AT, Taiwan; 
AJ, Japan; NA, North America. (Data resources: D’Abrera 1993, Inomata 1986, Makino 
1989, Shir6zu 1961, Walters, 1964, Zhang 1987.) 


Fagus Sibataniozephyrus Theclini 
EU Pe 0 3 
AC 6 0 58+ 
AT 1 1 25 
AJ 26 I 24 
NA ] 0 3 


yrus since that time (e.g., Kawazoé & Wakabayashi 1976, Saigusa 1983, 
Fukuda et al. 1984, D’Abrera 1993). 

In 1986 Inomata (1986) proposed the genus Sibataniozephyrus for 
the species fujisanus, but provided no phylogenetic analyses to support 
this placement. Indeed, no taxonomic level within the Theclini has been 
the subject of cladistic analysis to elucidate the phylogeny. 

Sibataniozephyrus represents the only Fagus-feeding member of 
Theclini. Because the distributional centers of both Fagus and Theclini 
are located in China (Table 1), where the Fagus-feeding behavior likely 
evolved, it seems unusual that Sibataniozephyrus was absent from there. 
Intrigued by this incongruity, we began extensive collecting in Fagus 
forests on Taiwan in 1986. These efforts finally produced an undescribed 
species of Sibataniozephyrus in 1992. 


MATERIALS AND METHODS 


A cladistic analysis of Sibataniozephyrus and its nearest relatives was 
conducted in an attempt to elucidate the relationship between Quer- 
cusia and Sibataniozephyrus. In addition to these two, the following 
genera were included in the cladistic analysis: Chrysozephyrus Shir6ézu 
& Yamamoto, Thermozephyrus Inomata, Neozephyrus Sibatani & Ito, 
Favonius Sibatani & Ito (divided into two OTU’s because it contains a 
remote member F. saphirinus Staudinger), and Austrozephyrus How- 
arth. Selection of these genera was based primarily upon studies by 
Shir6zu and Yamamoto (1956) and Eliot (1973). Two remote genera, 
Japonica Tutt and Euaspa Moore, were included in the analysis as 
outgroups. More than one outgroup was used because multiple out- 
groups will generate the most parsimonious cladogram globally (Mad- 
dison et al. 1984). Japonica was selected because its members retain a 
large number of ancestral characters (Shirézu & Yamamoto 1956). Euas- 
pa was chosen because the systematic position of Austrozephyrus was 
suspected to have common rooting with this genus (Saigusa 1983). 

Whenever possible, we used the type species of the above genera. 


130 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


When insufficient material was available for the type species, we used 
specimens of closely related species in the same genus. For selection of 
appropriate species and characters for the analysis, we consulted the 
following literature: Shir6zu and Yamamoto (1956), Shirézu (1960) and 
Shirézu and Saigusa (1970). Together, these publications contain de- 
tailed illustrations of numerous species of the genera examined in the 
cladistic analysis. All of the species substituted for the type species 
shared the same character states with the type species for all of the 
characters used in the analysis. Terminology for genitalic structures 
used in the character analyses and descriptions follow Klots (1970). 
Material examined (type species of each genus in bold): 

Chrysozephyrus disparatus (Howarth) (16, Taiwan); C. duma (Hewitson) (16, Nepal); C. 
kabrua (Tytler) (26, Taiwan); C. nishikaze (Araki & Sibatani) (16, Taiwan); C. rarasan- 
us (Matsumura) (64, Taiwan); C. smaragdinus (Bremer) (4619, Japan); C. souleanus 
(Riley) (2629, China); C. yuchingkinus Murayama & Shimonoya (26, Taiwan). 

Thermozephyrus ataxus (Doubleday & Hewitson) (3682, Japan; 1619, Taiwan). 

Neozephyrus japonicus (Murray) (86, Japan); N. helenae Howarth (19, China); N. tai- 
wanus (Wileman) (3612, Taiwan). 

Sibataniozephyrus fujisanus (Matsumura) (3689, Japan); S. kuafui Hsu & Lin (24619, 
Taiwan). 

Quercusia quercus (Linnaeus) (2622, France; 2619, Belgium). 

Favonius jezoensis (Matsumura) (6622, Japan); F. latifaciatus (Shir6zu) (2612, Japan); F. 
orientalis (Murray) (36, Japan); F. saphirinus (Staudinger) (4619, Japan); F. taxila 
(Bremer) (4¢, Japan). 

Austrozephyrus obsolon (Hewitson) (2642, Malaya). 

Japonica lutea (Hewitson) (16, Japan; 265°, Taiwan); J. saepestriata (Hewitson) (4689, 
Japan; 1612, China). 

Euaspa milionia (Hewitson) (26, Taiwan; 12, Nepal). 


The cladistic analysis was performed primarily using Hennig 86 
version 1.5 with implicit enumeration option (Farris 1988). Hennig 86 
does not allow multiple outgroup assignment, so only Japonica was 
assigned as outgroup, although Euaspa was retained in the analysis. 
Euaspa was expected to be linked with Austrozephyrus if they are more 
related to each other than to the rest of the ingroup members. PAUP 
version 2.4 also was employed to check the results, using global branch- 
ing swapping and multiple parsimony options (Swofford 1985). PAUP 
allows multiple assignments of outgroups, so both Japonica and Euaspa 
were regarded as outgroups in that analysis. 


Characters Used to Construct Hypothetical Phylogeny 
for Sibataniozephyrus and Its Related Genera 


Because of the uniform external features among these genera, gen- 
italic structures comprised most of the characters used in the analysis. 
Multiple state characters were used to construct the original data matrix. 
Hennig 86 allows two-way transformation series; the character coding 
in the data matrix (Table 2) and the character descriptions given below 


VOLUME 48, NUMBER 2 , 181 


TABLE 2. Data matrix. Column = characters, 0 = plesiomorphic state, 1, 2, 3, 1’, 2’ 
= derived states; row = genera: EUA, Euaspa (outgroup); JAP, Japonica (outgroup); CHR, 
Chrysozephyrus; THE, Thermozephyrus; NEO, Neozephyrus; SIB, Sibataniozephyrus; 
QUE, Quercusia; FAV, Favonius excluding F. saphirinus; FAS, Favonius saphirinus; 
AUS, Austrozephyrus. 


1 1 1 1 1 1 

1 2 3 4 5 6 1 8 9 0) 1 2 3 4 5 6 1 

JAP On me Ome? O00) 90-0 0; O “0° 0 0 0-0. °0 0 
Deen O OO .0O .0 0 .0 -0.1)..0 17:0, 0 ,1.0,..0 
ttwmmcmeerO eo 0. On 0. O 0. 06 O:.4d562 “Leek 0. 1 
ttm: O° 0: O° 0 OL O° °O “1 s.2--1°.1 0: | 
Cems Or Or OO. O20 ole 2.0) LO 
SIB OR le Ort Oere ls Oodle ily (On 200 6 O)> (0% 29) 0 Als, OF od 
twee FO 2° O 0 1 0. 1.40 /50°:0 1° 071. - 1 «21 
Poe 2  O 0 oO. 0: 2°°0 VY 0 221.1 0 1 
FAS CelemeOrers SOW S08 OF LO 22> 0 Oo Oo 20) Lome] (eek 
OMe teem O (Oo O60... 0... -0 8102 lv. 0) 7] 


are two-way. When PAUP was used, all the characters with two-way 
transformation series were divided into two characters with one-way 
transformation series. 

We polarized the characters mainly by outgroup comparisons. When 
the plesiomorphic state was not assigned to outgroups, explanations are 
followed by the character descriptions. In all character descriptions, 
the presumed plesiomorphic state (0) is listed first, followed by a trans- 
formation series in one direction (1, 2, ...) and a transformation series 
in another direction (1', 2’, .. .) respectively, when necessary. 

1. Brachium on tegumen: smooth and simple (Figs. 1, 24, 29) (0); 
smooth but flattened (Fig. 2) (1); serrated anteriorly (Fig. 3) (2); similar 
to state 2 but with additional large serrated lobe extending anteriorly 
(Fig. 4) (8); smooth, with basal half enlarged (Fig. 5) (1’); similar to 
state 1’ but bearing numerous minute teeth in front, (Fig. 6) (2’). 

2. Articulation of brachium with tegumen: double-articulated (0); 
single-articulated (1); brachium fused with tegumen, no articulation 
(2). 

3. Brachium general shape: hook shape (Figs. 1-6) (0); rod shape 
(Fig. 7) (1). 

4. Phallus structure: symmetrical with no special modification (Fig. 
8) (0); generally symmetrical but bearing an elongate dentate sclerite 
at the right side of aedeagus (Fig. 10) (1); similar to state 1 but dentate 
sclerite reduced with left side of aedeagus also reduced (Fig. 11) (2); 
caudal end of aedeagus slightly asymmetrical, weakly sclerotized (Fig. 
12) (1’); caudal end of aedeagus strongly asymmetrical, forming com- 
plicated, heavily sclerotized structure (Fig. 13) (2’).. 

The character state (1’) of Japonica was considered derived because 


132 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Le 
: 


br an 


ees 


Fics. 1-6. Left brachium: 1, Japonica saepestriata; 2, Neozephyrus japonicus; 3, 
Chrysozephyrus smaragdinus; 4, Thermozephyrus ataxus; 5, Favonius orientalis; 6, 
Quercusia quercus (scale line = 1 mm). 

Fic. 7. Lateral view of sclerites of 9 + 10 genitalic segments with modified left 
brachium attached in Austrozephyrus obsolon; br, brachium, un, so-called “uncus” with 
its homology with the other uncus-like structures found in the other Theclini uncertain 
(scale line = 1 mm). 


VOLUME 48, NUMBER 2 JLB38} 


the simple, symmetrical phallus (0) is found commonly in ingroup 
genera and also in the other outgroup genus Euaspa. 

5. Phallus upcurved: no (Fig. 9) (0); yes (Figs. 25, 30) (1). 

6. Aedeagus twisted: no (Figs. 8, 10-18) (0); aedeagus twisted 90° 
toward the left side (Fig. 14) (1). 

7. Juxta shape: U- or V-shaped (Figs. 15, 16) (0); modified into a 
simple plate (Figs. 17, 26, 31) (1). 

8. Harpal region of valva: no process (Figs. 18-21) (0); bearing a 
prominent elongate process proximately (Figs. 27, 32) (1). 

9. Ampulla of valva: not serrated (Figs. 18, 27, 32) (0); finely serrated 
mesad (Fig. 19) (1); strongly serrated mesad (Fig. 20) (2); modified into 
a large erect, dentate tooth (Figs. 21, 22) (1’). 

10. X tergite: uncus absent (Figs. 28, 28) (0); so-called ‘‘uncus’’ (with 
homology uncertain) in various forms present (Fig. 7) (1). 

The plesiomorphic state was not assigned to the so-called ‘“‘uncus”’ 
found in Euaspa because this uncus-like structure is found only in 
Austrozephyrus in the ingroup and its homology with Euaspa was 
uncertain. This decision is supported by the fact that both Ogyrini and 
Arhopalini, the most likely sister groups of Theclini, lack an uncus (Eliot 
1973). 

11. Two signa on corpus bursa: small, rounded, invaginated inwards 
into a spine (Figs. 35, 36, 38) (0); extended enormously into a spiny, 
elongate stripe (Fig. 34) (1); lost (Fig. 38) (1’). 

12. Lamella postvaginalis of sterigma: bilobed, separated into two 
pieces (Figs. 33, 36, 38) (0); modified into a single rectangular plate, 
sometimes with complicated structures (Fig. 34) (1); modified into a 
double layered plate, with the ventral layer elongate and rectangular 
while the dorsal layer bifid, finger-like, and projecting posteriorly (Fig. 
35) (1’). 

The character state (1') of outgroup Euaspa is apparently not ple- 
siomorphic since this state is specific to Euaspa and not found anywhere 
else in Theclini. 

13. Sexual dichromatism: absent (0); present, male with specialized 
purple metallic scales on upperside of the wings, female dull-colored 
(1); similar to state 1 but with green/blue metallic scales (2). 

14. Female polymorphism: absent (0); present, individual with blue 
patch, orange spots, or both present or absent on forewing upperside 
(aly). 

15. Foretarsus: segmented in both sexes (0); rudimentary in male (1). 

16. Forewing venation: M1 forked with R8 away from the base (0); 
M1 forked with R3 at the base (1). 

17. Male eyes: naked or sparsely hairy (0); densely hairy (1). 


134 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Lo 


—— er 


Fics. 8-14. Phallus: 8, Chrysozephyrus smaragdinus (dorsal view); 9, Chrysozephy- 
rus smaragdinus (ventral view); 10, Neozephyrus japonicus; 11, Favonius orientalis; 
12, Japonica saepestriata; 13, Quercusia quercus; 14, Austrozephyrus obsolon (scale 
line = 1 mm). 


VOLUME 48, NUMBER 2 135 


7 


Fics. 15-17. Juxta: 15, Japonica saepestriata; 16, Chrysozephyrus smaragdinus; 17, 
Quercusia quercus (scale line = 1 mm). 

Fics. 18-22. Right valva: 18, Chrysozephyrus smaragdinus; 19, Quercusia quercus; 
20, Favonius orientalis; 21, Thermozephyrus ataxus; 22, Lateral view of right ampulla 
of Thermozephyrus ataxus (scale line = 1 mm). 


136 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


mew UL Le ew mm mw ew ew em wm eer wm mm em ee 


Fics. 23-27. Male genitalia of Sibataniozephyrus fujisanus (Matsumura): 23, Lateral 
view of sclerites of 9 + 10 genitalic segments with brachia and left valva attached; 24, 
left brachium; 25, phallus; 26, juxta; 27, dorsal view of right valva. 

Fics. 28-32. Male genitalia of Sibataniozephyrus kuafui, new species: 28, Lateral 
view of sclerites of 9 + 10 genitalic segments with brachia and left valva attached; 29, 
left brachium; 30, phallus; 31, juxta; 32, dorsal view of right valva. . 


VOLUME 48, NUMBER 2 137 


Fics. 33-35. Female genitalia: 33, Favonius jezoensis; 34, Neozephyrus taiwanus; 
35, Euaspa milionia (scale line = 1 mm). 


RESULTS OF THE CLADISTIC ANALYSES 


Two minimum length trees (Figs. 40, 41) were derived from the data 
set using Hennig 86 each with CI = 0.78. Two equally parsimonious 
trees of the same topologies as above were found using PAUP each 
with CI = 0.81. The ingroup topology did not change when the out- 
groups were removed. 

The high CI values of the trees (CI = 0.78/0.81) indicate that the 
data matrix has relatively little homoplasy. 

Austrozephyrus, considered by Saigusa (1983) to be closely related 
to Euaspa, forms a monophyletic clade with Quercusia lineage distinct 
from Euaspa, supported by characters 2, 13, and 17 (Figs. 40, 41). Thus 
it should be retained within the ingroup. Whether Sibataniozephyrus 
should be treated as congeneric with Quercusia is still somewhat dis- 
putable. Of the two equally parsimonious trees produced, one suggests 
a monophyletic relationship between Sibataniozephyrus and Quercusia 
(Fig. 40). Nevertheless this tree seems less likely to reflect the true 
phylogeny because the only synapomorphy for the two genera is a 
platelike juxta (character 7). With the rest of the ingroup and outgroup 
genera possessing a V- or U-shaped juxta, the platelike juxta found in 
Sibataniozephyrus and Quercusia may reflect the loss of function owing 
to the specialized phallus of the two genera. The strongly upcurved 
phallus of Sibataniozephyrus and the elongate phallobase and enlarged 


138 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


1mm 


Fics. 36-37. Female genitalia of Sibataniozephyrus fujisanus (Matsumura): 36, Cor- 
pus bursa with stigma; 37, papilla analis with apophysis posterioris. 

Fics. 38-89. Female genitalia of Sibataniozephyrus kuafui, new species: 38, Corpus 
bursa with stigma; 39, papilla analis with apophysis posterioris. 


aedeagus of Quercusia probably function to stabilize the phallus in the 
male genitalic complex. Thus it is our view that the platelike juxta can 
not be regarded unequivocally as a synapomorphy for these two genera. 
Furthermore, this tree suggests the most derived state of character 13 
(i.e., metallic blue/green color in male) evolved twice: once on the 
lineage of Sibataniozephyrus and the other on the lineage leading to 


VOLUME 48, NUMBER 2 139 


~ 0 
2) 
~ S w 2 
< & ES 
Q rm) > 7) = £ 
7) 3 < 2 > Q 
w q = o > 6 2 
45] O — A 2) N £ CT) q 
AS) wo | c < 2 fe) Q N 
c Q (s) & Q ¢ Ss © fe) = 
S a o 8 3 S % 3 2 o 
=} = . ® <= < 
S Ar 9g GH u Ar < = os) - 
12-3 
1-2 0-1 
2 f ae gol 
19-2" 501 169-1 10-1 ayia 90-1 
1 32 
119-1 a 
60-1 
0 1 
10 40 —2' 80-1 
0 1 
402 10"? 10-1 
0>1' 0>1' 
4 = 12°> 169-1 131-2 
Q0 2 
120-1 
119>1' 
131-2 
0-51 
ag 140-1 
20-1 
139-1 
17°! 
159-1 


Fic. 40. A cladogram less likely to reflect the true phylogeny of Sibataniozephyrus 
and possibly related genera (produced primarily by Hennig 86 but with slight modification 
at the base since character 15 is more logical to have segmented tarsus assigned as 
plesiomorphic state). 


140 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


WwW 
2 7) “” 
2 2 g c 
o S 2 5 = < 
N r= q = = 
& [e) — i‘b) > iS) 
% a 2 aS 7) N = Q N 
Q = iS < =~ Ss} ro Q 9 
= & o roy NN N 
cf BA Bee 8, 82 ee Ee ee 
Q © S Q - > 3 9 re @ 
s a CS) oj u hg < 2 S Fe 
12-3 
1-2 0-1 
2 ah go 1 
19-2’ | 50-1 1609-1 191 90-1 
30-1 : 
{ 11-2 
: 110-1 
1001 60-1 
40 —2' 70-1 
0-1 
40 2 LOST 10-1 
0>1' 0>1' 
Ay 12 yo! 80-1 Q0 2 
0-1 
119-1 12°> 
16°! 
140-1 
131-2 
20-1 
130-1 
1701 
150-1 | 


Fic. 41. Proposed phylogeny based upon the preferred cladogram for Sibataniozeph- 
yrus and possibly related genera (produced primarily by Hennig 86 but with slight 
modification at the base since character 15 is more logical to have segmented tarsus 
assigned as plesiomorphic state). 


VOLUME 48, NUMBER 2 141 


the rest of the metallic green/blue-winged genera. However, because 
Sibataniozephyrus and the majority of the other blue/green-winged 
genera are found in the same geographical range in Asia and not found 
in the other continents where Theclini also are present, the metallic 
green/blue scales probably evolved only once. The cladogram shown 
on Fig. 41 shows a paraphyletic relationship between Quercusia and 
Sibataniozephyrus. Both genera have a few autapomorphies which 
justify their separation as genera in the annotated Linnaean classifi- 
cation conventionally used. Quercusia possesses characters 1, 4, 16 and 
Sibataniozephyrus characters 5 and 7. 

It is our view that Sibataniozephyrus should be recognized as an 
independent genus and the superficially similar platelike juxta is ho- 
moplastic. Based on the preferred proposed phylogeny, the purple 
Quercusia is a sister group of all the other genera having metallic green 
or blue scales in the male. All these green/blue-winged genera are 
found only in east and southeast Asia; Quercusia and the green/blue- 
winged genera are allopatric. It seems that Quercusia diverged early 
in the evolutionary history of these groups and extensive evolutionary 
change took place between the two lineages. Sibataniozephyrus also 
diverged early and underwent significant specialization as it obtained 
its unique niche of Fagus-feeding, a feature found nowhere else in 
Theclini. These specializations probably occurred after the lineage of 
the common ancestor of the green/blue-winged genera already split 
from the lineage of the ancestor of Quercusia, which feeds on a wide 
range of plants in different families but is not known to feed on Eu- 
ropean Fagus (Shirézu 1961, Carter 1982). It is apparent that symple- 
siomorphic characters shared by these two lineages of Theclini led to 
Shir6zu and Yamamoto’s (1956) decision to treat them as congeneric. 


TAXONOMY 
Sibataniozephyrus Inomata, 1986 


Sibataniozephyrus Inomata 1986:120. 


Type-species. Zephyrus fujisanus Matsumura, 1910, by monotypy. Gender masculine. 

Diagnosis. Moderate sized, diurnal butterflies (FW length 15.0-18.6 mm). Head: Hairy, 
clothed with upraised, black and grayish white hairs on vertex and frons, appressed scales 
on occiput, a white rim surrounding the eye; eye semi-oval, hairy; labial palpus hairy, 
porrect, pointed, projecting ahead of plane of front; maxillary palpus reduced, not visible; 
proboscis unscaled; antenna smooth-scaled, with projecting setae at nudum in both sexes, 
antennal length in male longer than in female. Thorax: Grayish white dorsally, tinged 
with metallic structural colors under reflection of light in male, uniformly brown in 
female; white ventrally, legs white, banded with dark brown on tarsi. Abdomen: Dark 
brown dorsally, white ventrally. Forewing: Termen nearly straight, costa curved; 11 veins, 
R4 + 5 and M1 both forked with R38, the forking points dividing R8 into three nearly 
equal long parts; upperside scaling shining metallic with a dark dull margin in male, 
uniformly dark in female, both sexes with underside markings visible by transparency; 
underside scaling white to pale brown with series of dark submarginal spots between two 


142 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


transverse lines; a prominent dark discal band, a white patch between this band and the 
submarginal spot system regardless of ground color. Fringe white. Hindwing: Slender 
“tail” from Cu2; 9 veins all separate; upperside scaling shining metallic with a dark 
margin always broader than that of forewing in male, uniformly dark in female, both 
sexes with a very fine white outline along termen, underside markings visible by trans- 
parency; underside scaling similar to that of forewing but with a large yellow or orange 
tornal patch with black rounded spot in Cul, and a black irregularly shaped spot at the 
tornus, white outline also present; “‘tail’’ black with white tip on both surfaces. Fringe 
generally white, but outer fringe around tornus dark brown. Male genitalia: Sclerites of 
9th and 10th segments fused into a complete ring, with tegumen of two segments indis- 
tinguishable; uncus absent; socii folded deeply inwards; brachium single-articulated with 
tegumen, smooth, hook-like; saccus fairly short; phallus symmetrical, strongly upcurved; 
juxta platelike; valva semicircular, ampulla forming a long spine, harpal region bearing 
a slender process proximally, fused with the lobelike cucullus + valvula distally, costa 
prominent and elongate, sacullus relatively well developed with a solid ventral ridge. 
Female genitalia: Apophyses posteriores elongate, slender, with weakly sclerotized, some- 
what square-shaped papillae anales; ductus bursa slender, heavily sclerotized; sterigma 
with lamella antevaginalis absent, lamella postvaginalis heavily sclerotized, forming a 
pair of spines projecting posteriorly; corpus bursa oval, bearing a pair of small, invaginated 
spinelike signa. 


Biology of Sibataniozephyrus 


The biology of S. fujisanus has been described in detail by Shirézu 
and Hara (1960) and Fukuda et al. (1984). It will be briefly summarized 
below because both works are in Japanese. The exclusive host plants of 
S. fujisanus are Fagus crenata Blume and F. japonica Maximowiczi. 
Ovum is white and fairly large (diameter 1.00-1.02 mm, height 0.54 
mm). It is laid singly on a twig and hibernates. The larva hatches in 
early spring when the host starts budding, and initially bores into buds. 
The fully grown larva is yellowish brown, reaching 16.5 mm. It makes 
a shelter by spinning two or three leaves together, and consumes young 
leaves and buds. Feeding usually takes place nocturnally. The pupa is 
uniform light brown with two rows of subdorsal dark dots. Pupation 
usually takes place under fallen leaves around the base of the host. 
Sibataniozephyrus kuafui, the new species described below, presum- 
ably feeds on F. hayatae Palibin ex Hayata because this butterfly is 
associated exclusively with this plant. 


Sibataniozephyrus fujisanus (Matsumura) 
(Figs. 28-27, 36, 37, 42-45) 


Zephyrus fujisanus Matsumura 1910:221. 

Zephyrus nohirae Matsumura 1915:1, pl. 1, fig. B. 

Zephyrus attilia ab. subgriseus Wileman 1911:55. 

Zephyrus fujisanus ab. yamamotoi Uchida 1932:232, tab. 16, fig. 5. 

Zephyrus fujisanus ab. zoasanus Kanda 1933:8. 

Favonius fujisanus; Sibatani & Ito 1942:327. 

Quercusia fujisanus; Shir6zu & Yamamoto 1956:393. 

Ouercusia fujisanus latimarginatus Murayama 1963:48, figs. 31, 33, 35, 37. syn. n. 
Sibataniozephyrus fujisanus; Inomata 1986:120 (synonymy). 


VOLUME 48, NUMBER 2 143 


TL. Mt. Fuji [Honshu, JAPAN]. 

Diagnosis. Male: Forewing length 16.0-18.0 mm (mean = 17.0 mm, n = 8); antennal 
length 8.0-8.8 mm (mean = 8.35 mm, n = 3). Forewing upperside bright metallic bluish 
green, margin dark brown; underside ground color yellowish white or white tinged with 
brown, all markings brown with a very stout transverse line adjacent and proximal to the 
submarginal spots. Hindwing upperside with coloration similar to that of forewing but 
with a much broader dark brown margin, its narrowest part still wider than the broadest 
part of the forewing margin, the white outline weakly developed; underside with the 
same ground color as forewing, discal band and discoidal bar separate, tornal patch yellow. 
Male genitalia: Ring-shaped sclerites of 9 + 10 segments width 0.56 times height; tegumen 
9 + 10 forming a bump dorsally; brachium twisted; phallus with a short, stout aedeagus, 
length 1.1 times phallobase; valva with a short spine on ampulla, 0.2 times as long as 
entire valva, distal margin of the harpal region and ventral ridge of sacculus smooth, not 
serrated. 

Female: FW length 15.0-17.2 mm (mean = 16.5 mm, n = 8); antennal length 7.0-7.8 
mm (mean = 7.2 mm, n = 8). Both wings with upperside ground color uniformly brown, 
underside pale brown with an orange tornal patch, otherwise similar to male. Female 
genitalia: Lamella postvaginalis spines long and slender, each with a semicircular lateral 
lobe near the base, arranged nearly parallel to each other, with distance between the tip 
of the two spines approximately 0.19 mm; distance from the tip of the spine to ostium 
0.53 mm; ostium width 0.01 mm. 

Species examined. JAPAN: 29, Honshu, Tottori Pref., Mt. Daisen, 25. VI. 1973 (T. 
Yuzuriha); 16192, same locality as above, 25. VI. 1988 (Y. Hama); 1622, Honshu, Kyoto 
Pref., Mt. Oe, emgd. 11. V., 15. V. and 27. V. 1988 (S. Kugo); 12, Honshu, Kyoto Pref., 
Tamba Town, Funai-gun, emgd. 10. V. 1977 (T. Yuzuriha); 12, Honshu, Kyoto Pref., 
Asauegun, Yamakawamachi, Mt. Kouetsu, emgd. 14. V. 1984 (T. Yuzuriha); 146, Yamanashi 
Pref., Tomurozawa, emgd. 15. V. 1984 (T. Yuzuriha); 12, Hokkaido, Kaminokunicho, 
Zentana, emgd. 8. VI. 1986 (T. Doi). 

Geographical distribution. S. fujisanus is endemic to Japan (southern Hokkaido, Hon- 
shu, Shikoku, and northern to central Kyushu). 

Comments. The populations in Kyushu, southern Japan, have been given subspecific 
status based upon two characters: the broader dark margin on upperside of the wings 
and the darker color on the underside of the wings. However, Inomata’s (1986) discussion 
and numerous pictures of samples from various regions in Japan clearly reveal that 
variation within the two traits is continuous. The southern populations have a stronger 
tendency to show the two traits, but specimens indistinguishable from those from central 
Japan are encountered. This is clearly an example of clinal variation and in our opinion 
should not be recognized nomenclaturally. 


Sibataniozephyrus kuafui Hsu & Lin, new species 


(Figs. 28-32, 38, 39, 46-49) 


Description. Male: Forewing length 16.1-18.6 mm (mean = 17.08, n = 24); antennal 
length 8.2-9.0 mm (mean = 8.5 mm, n = 24). Forewing upperside metallic dark blue 
tinged with green, margin black; underside ground color white with submarginal spots 
dark black, other markings dark gray, the proximal transverse line very slim. Hindwing 
upperside with the metallic area greenish blue with a purplish outline, margin black, not 
as broad as in S. fujisanus, the termenal outline distinct, forming a white line through 
entire termen; underside with the same ground color as of the forewing, discal band and 
discoidal bar always completely merged, not distinguishable from each other, tornal patch 
orange or yellowish orange. Male genitalia: Ring-shaped sclerites of 9 + 10 segments 
width 0.65 times height; tegumen 9 + 10 with dorsal side flat; brachium forming a simple 
hook, not twisted as in S. fujisanus; phallus with a long and slender aedeagus, length 1.4 
times phallobase; valva with a long spine on ampulla, 0.3 times as long as entire valva, 
distal margin of the harpal region bearing 4 to 10 teeth (n = 9), number not always equal 
on opposite valvae, ventral ridge of sacculus heavily serrated. 


144 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 42-45. Sibataniozephyrus fujisanus (Matsumura) (JAPAN, Honshu): 42, 4, 
upperside; 43, 6, underside; 44, 2, upperside; 45, 2, underside (scale line = 1 cm). 


Female: Forewing length 16.0 mm (n = 1); antennal length 7.0 mm (n = 1). Both 
wings with upperside ground color dark brown and underside markings more developed 
than those in the male, otherwise similar to the male. Female genitalia: Lamella post- 
vaginalis spines short and stout, no lateral lobe; the two spines widely separated, diverging, 
distance between tips of the two spines approximately 0.46 mm; distance from the tip of 
the spine to ostium 0.30 mm; ostium width 0.01 mm. 

Type material. Holotype 6, TAIWAN: Taipei Hsien, Sanhsia, Mt. N. Chatienshan, 
1700-1727 m. 121°27’E, 24°47.5'’N, 22. V. 1993 (Y. F. Hsu & M. Y. Lin) (NTUIM). 
Paratypes. 9612, same data as holotype (12 dissected: Genitalia YFH 0747); 146, same 
locality as holotype, 28/29. V. 1992 (98 dissected: Genitalia YFH 0441, 0442, 0447, 0487, 
0498, 0499, 0500, 0501, 0502) (Y. F. Hsu & M. Y. Lin) (BMNH; CAS; IZASB; NTUIM; 
UCB; NMNH). 

Depositories. Abbreviations of institutions are as follows: the Natural History Museum, 
London (BMNH); California Academy of Sciences, San Francisco (CAS); Zoological In- 
stitute of Academia Sinica, Beijing (IZASB); the Insect Museum, National Taiwan Uni- 
versity, Taipei (NTUIM), Essig Museum of Entomology, University of California, Berkeley 
(UCB), and U.S. National Museum of Natural History, Washington, D.C. (NMNH). 

Geographical distribution. S. kuafui is known from Taiwan. 

Etymology. Kuafu is a character in Chinese mythology who tried to challenge the sun 
by chasing it until his exhausting death. 

Comments. Sibataniozephyrus kuafui undoubtedly utilizes Fagus hayatae as its larval 
host in Taiwan because it is associated exclusively with this plant. Fagus hayatae, formerly 


VOLUME 48, NUMBER 2 j 145 


Fics. 46-49. Sibataniozephyrus kuafui, new species (TAIWAN): 46, 6, upperside; 
47, 6, underside; 48, 2, upperside; 49, 2, underside (scale line = 1 cm). 


recognized as Taiwan endemic, recently has been reported from central China and was 
noted by Shen and Boufford (1988) as a remarkable new example of disjunction between 
Taiwan and central China. This discovery suggests that the population of S. kuafui in 
Taiwan may be a relict and suggests the possibility of finding this insect on mainland 
China. Shen and Boufford (1988) also point out that F. lucida and F. hayatae are more 
closely related to each other than they are to any other extant species of Fagus. According 
to Tsien et al. (1975:11), F. lucida is distributed approximately between 24°N and 30°N, 
which covers the latitudinal range where F. hayatae and S. kuafui are found in Taiwan. 
This fact makes F. lucida another candidate as a larval host for S. kuafui, or a possible 
sister species, on the Chinese mainland because this beech is apparently far more abundant 
than F. hayatae there. 


CONCLUSION AND DISCUSSION 


We conclude that Sibataniozephyrus should be considered a valid 
genus distinct from Quercusia on the basis of their phylogenetic rela- 
tionship, biology, and biogeography. Each of these two genera retains 
several symplesiomorphic characters, while each has gained autapo- 
morphic features. This phenomenon led early workers to lump them 
into a single genus. 


146 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


We predict that Sibataniozephyrus will eventually be found on main- 
land China because China supports the richest diversity of both Fagus 
and Theclini in the world (Table 1). Moreover, it seems likely that more 
than one Sibataniozephyrus species may be present on mainland China 
based on the following observations: 

1) There are multiple species of Fagus in China, and these species 
show various degrees of allopatry. Even when multiple species are 
sympatric, each species usually is dominant in a different topogeo- 
graphical and horizontal zone (Tsien et al. 1975), which might favor 
vicariant speciation because relictual Fagus populations may function 
as isolated ecological and evolutionary islands. 

2) Pollen studies reveal that Fagus species lived in many areas where 
they are absent today (Steenis 1971) indicating Fagus populations have 
suffered extensive extinctions in the past, which also may have led to 
vicariant speciation. 


ACKNOWLEDGMENTS 


We thank Jerry A. Powell (UCB) and John T. Doyen (UCB) for helpful discussions on 
the cladistic analyses and for critically reading the manuscript. We also thank Hideyuki 
Chiba (JAPS), Chang-Fu Hsieh (NTU), Chun-Lin Li (NTU) and Toshio Inomata (Japan) 
for literature searching, Scott A. Stockwell (U.S. Navy) for helping with the use of the 
Hennig 86 software, and Takashi Yuzuriha (Japan) and Seiho Kugo (Japan) for offering 
the specimens from Japan. 


LITERATURE CITED 


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Fukuba, H., E. Hama, T. Kuzuya, A. TAKAHASHI, M. TAKAHASHI, B.. TANAKA, H. 
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SHEN, C. F. & D. E. BOUFFORD. 1988. Fagus hayatae (Fagacae)—A remarkable new 
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4:231-233. 

WALTERS, S. M. 1964. Fagus. In Tutin, T. G., V. H. Heywood, N. A. Burges, D. H. 
Valentine, S. M. Walters & D. A. Webb (eds.), Flora Europaea: 61. Cambridge 
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Sinicorum, Supplementum I. Science Publishing Ltd., Beijing. [in Chinese]. 


Received for publication 20 October 1993; revised and accepted 23 January 1994. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 148-156 


AVIAN PREDATION ON THE MONARCH BUTTERFLY, 
DANAUS PLEXIPPUS (NYMPHALIDAE: DANAINAE), AT A 
CALIFORNIA OVERWINTERING SITE 


WALTER H. SAKAI! 


Life Science Department, Santa Monica College, 
1900 Pico Boulevard, Santa Monica, California 90405-1628, USA 


ABSTRACT. Avian predation on monarch butterflies (Danaus plexippus L.) was 
observed at a California monarch overwintering site. A pair of rufous-sided towhees 
(Pipilo erythrophthalmus Linneaus) exerted a 6.51-7.43% annual mortality on the over- 
wintering colony, consuming an average of 22.7 butterflies/day. A characteristic predatory 
signature was observed on the uneaten remains left by the towhees. Tagging studies 
indicate that tagged butterflies were at greater risk of predation than were untagged 
butterflies. 


Additional key words: rufous-sided towhee, sex ratios, cardenolides, tagging. 


There are several reported examples of predation of monarch but- 
terflies at overwintering sites. In Mexico, avian predators include black- 
headed grosbeaks (Pheuctius melanocephalus Swainson), Scott’s oriole 
(Icterus parisorum Bonaparte), and black-backed oriole (Icterus abeillei 
Lesson) (Calvert et al. 1979, Brower & Calvert 1985). In California, 
avian predators include chestnut-backed chickadees (Parus rufescens 
Townsend) (Tuskes & Brower 1978, Brower & Fink 1985, Bell & Dayton 
1986), starlings (Sturnus vulgaris L.) (Dayton & Bell, pers. comm.), and 
scrub jays (Aphelocoma coerulescens Bosc) (Peterson, pers. comm.). 
Mammalian predators found at Mexican overwintering sites include 
Peromyscus spicilegus J. A. Allen, Microtus mexicanus salvus Hall, 
and Peromyscus melanotis (=P. maniculatus labecula Elliott) (Brower 
et al. 1985, Glendinning et al. 1988). Leong et al. (1990) reported 
monarch predation in California by yellowjackets (Vespula vulgaris). 
In the 1989-90 season, I discovered a pair of rufous-sided towhees 
(Pipilo erythrophthalmus Linneaus, Emberizidae) preying upon mon- 
archs in a moderate-sized overwintering colony in the Santa Monica 
Mountains, Los Angeles County, California. During the course of tag- 
ging monarch butterflies at this site, | measured predation rates. 


METHODS 


From 13 September 1989 to 28 March 1990, I made weekly visits to 
a monarch overwintering site along the coast in Los Angeles County, 
California, 18 km west of Malibu, to tag monarch butterflies. Tags of 
the type developed by Urquhart (1960, 1976, 1987) were used to de- 


1 Research Associate, Entomology Section, Museum of Natural History of Los Angeles County, 900 Exposition Blvd., 
Los Angeles, California 90007, U.S.A. 


VOLUME 48, NUMBER 2 149 


TABLE 1. Observations of a single pair of rufous-sided towhees feeding activities on 
31 January 1990. 


Time Activities observed 
06:53 h Sunrise (Anon 1989). 
07:00 h Unsuccessful sally. 
07:03 h Unsuccessful sally. Monarch flies, 
bird sally-strikes unsuccessfully. 
07:08 h Successful sally. 
07:13 h Successful sally. 
07:45 h Sunlight hitting site. 
No bird activity. 
08:03 h Picks up monarch fallen on ground. 
08:07 h Picks up monarch fallen on ground. 
08:13 h Successful sally. 
08:15 h Picks up monarch fallen on ground. 
08:27 h Picks up monarch fallen on ground. 
08:30 h Successful sally, does not eat it. 
08:31 h Successful sally. 
08:51 h Successful sally. 
09:01 h Successful sally. 


No bird activity after this. 


termine predation rate and selectivity. All butterfly remains were col- 
lected from the litter under the clustering areas during my weekly 
visits. On 12 and 13 January 1990, the site was visited, and butterfly 
remains were collected to measure daily predation rate. 

Field observations of bird predation were made by binoculars. Con- 
tinuous early morning observations from before sunrise until no further 
predation activity was observed were made on three occasions. Because 
rufous-sided towhees are not sexually dimorphic, I was unable to dis- 
tinguish between the sexes. Terms used to describe monarch predation 
by towhees follow Remsen and Robinson (1990). 


RESULTS 


Towhee activity recorded on 31 January 1990 is presented in Table 
1. The pair of towhees “‘perched” on small shrubs. Looking about, the 
towhee would “‘sally” up some 10-15 m into the tree, “pounce” upon 
a butterfly, and bring it to the ground near or under a shrub. On other 
occasions, the attacked butterfly fell to the ground, where the bird 
would pick it up. The towhee also attacked monarchs that previously 
had fallen to the ground. It appears that the towhee “‘snips”’ the butterfly 
between the head and thorax based on examination of intact dead 
butterflies. All four wings were removed by the bird by “‘snipping”’ 
between the body and wing and “shaking” the prey until the wings 
fell off before the body was consumed. Characteristic piles of the four 
wings were often found under or near shrubbery. 


150 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


TABLE 2. Tagging and recapture summary KSP site in 1989-90. 


é Q Total Recap Total 
Date tagged tagged tagged % } Recaptures % 6 captured 
4 Oct 89 3 1 4 75.0 0 0 4 
11 Oct 89 22 1 23 S15) Of 1 100 24 
18 Oct 89 63 41 104 60.6 4 100 108 
25 Oct 89 180 124 304 509.2 32 71.9 336 
1 Nov 89 157 124 281 00.9 itl 45.5 292 
13 Nov 89 242 219 461 52.5 15 60.0 476 
15 Nov 89 173 204 377 45.9 o7 61.4 434 
22 Nov 89 249 225 474 52.9 36 61.1 510 
29 Nov 89 135 115 250 54.0 30 70.0 280 
6 Dec 89 58 50 108 03.7 IZ 64.7 125 
13 Dec 89 275 265 540 00.9 67 65.7 607 
20 Dec 89 384 385 769 49.9 93 66.7 862 
27 Dec 89 362 330 692 52.3 90 61.1 782 
3 Jan 90 142 113 255 Do.7 80 58.8 335 
10 Jan 90 168 138 306 54.9 92 56.5 398 
17 Jan 90 Pare 169 440 61.6 183 57.9 623 
24 Jan 90 104 16 120 86.7 42 85.7 162 
31 Jan 90 267 14 281 95.0 67 95.5 348 
3 Feb 90 89 14 103 86.4 UZ 94.4 175 
10 Feb 90 84 2 86 O7.7 45 95.6 131 
14 Feb 90 47 4 ol 9222 30 96.7 81 
21 Feb 90 11 0 11 100 3 66.7 14 
28 Feb 90 2 0 2 100 0 0 2 
Totals 3488 2554 6042 57.7 1067 69.3 7109 


$= 6 tagged/(é tagged + 2 tagged); Recap % 6 = % of 6 recaptures; Total captured = total tagged + recaptures. 


Fresh intact but damaged butterflies also were found. Two were 
found walking around in the litter almost decapitated, damage only in 
the area between the head and thorax, where the towhee immobilized 
the prey by biting the butterfly. The heads, occasionally found in the 
litter, were then lost as the towhee removed the wings. 

Predation by towhees occurred in the early morning hours from an 
hour before until two hours after sunrise (see Table 1). The towhees 
were observed sallying (“‘sally-strike”) for flying monarchs on two oc- 
casions but were unsuccessful. On one occasion, a butterfly was dis- 
lodged from the tree and “‘flutter-chased” unsuccessfully. The sallying 
behavior occurred when the butterflies were beginning to fly and were 
quite slow. Predation was never observed during the rest of the daylight 
hours or during sunset, even though monarchs were seen roosting and 
flying about. 

Table 2 shows the tagging record for 6042 monarchs during the 1989- 
90 season at this site. The sex ratio is skewed favoring males (57.7%, 
Chi-square = 72.625, df = 1, P < 0.001). 

Predation was noticed first on 3 January 1990, rose dramatically 
thereafter, and remained high until late February when the colony size 


VOLUME 48, NUMBER 2 . 151 


TABLE 3. Summary of predated butterflies collected at weekly intervals from the 
litter at KSP site for 1989-90. 


Date fo) Q %s 


? Total x/day 
3 Jan 90 3 3 50.0 0 6 0.9 
10 Jan 90 29 Zit 51.8 2 58 8.3 
17 Jan 90 105 69 60.3 + 178 25.4 
24 Jan 90 81 58 58.3 i 146 20.9 
31 Jan 90 73 Dal 73.0 2 102 14.6 
3 Feb 90 74 56 56.9 0 130 43.6 
10 Feb 90 98 46 68.1 2Z 146 20.9 
14 Feb 90 133 A7 73.9 1 181 45.2 
21 Feb 90 163 66 71.2 2 231 33.0 
28 Feb 90 47 14 77.0 1 62 8.9 
7 Mar 90 22 9 dale) il 32 4.6 
14 Mar 90 10 6 62.5 0 16 2.3 
21 Mar 90 13 il 92.5 0 14 2.0 
Totals 851 429 66.5 22 1302 


m a sex undetermined; Total = 6 + 2 + ?; % 6 = 6/(6 + 9); x/day = average number of butterflies eaten per day each 
week. 


declined to a few hundred individuals. Predation ceased by 21 March 
1990 when the butterflies left. Table 3 shows the total number and sex 
ratio of butterflies remains collected at each sampling period. The rate 
of predation varied from 14.6 to 43.8 butterflies per day for samples 
from 17 January to 21 February 1990. Earlier samples were omitted 
as the towhees were learning to exploit the resource, and later samples 
were omitted as the resource diminished. The average for this period 
was 22.7 butterflies per day. Since this was the work of a single pair of 
rufous-sided towhees, each bird was eating an average of 11.4 butter- 
flies/day. 

Thirty-eight butterflies were found on 12 January, two days after the 
previous census, and 38 butterflies were found on 18 January. An av- 
erage of 19 butterflies was eaten on 10-12 January and 38 butterflies 
were eaten on 12-13 January. 

The total number of dead butterflies collected in the litter was 1302. 
Taking this colony with a peak population visually estimated at 20,000, 
this was a 6.51% mortality. A mortality estimate of 7.43% can be derived 
based on the number of butterflies tagged and the number of butterflies 
preyed upon by the towhees. These estimates were, however, misleading 
as the population size was steadily declining (visually estimated at 5000 
on 17 January) when predation plateaued. By late February, the towhees 
were taking most of the remaining butterflies; thus, the 6.51-7.43% 
annual mortality estimate is extremely conservative. 

A Chi-square was calculated comparing the actual sex ratio of the 
butterflies when the birds were feeding based on Table 2 with the sex 
ratio of eaten butterflies in Table 3. The Chi-square value of 0.185 (df 


152 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


TABLE 4. Summary of preyed upon tagged and untagged butterflies collected at 
weekly intervals from the litter at KSP site for 1989-90. 


Untagged Tagged 
Date 8 Q % 3 Xu $ Q % 6 X i T 
3 Jan 90 3 3 50.0 6 0 (0) Oo) 0) 6 
10 Jan 90 14 21 40.0 35 15 6 71.4 21 58 


17 Jan 90 76 58 56.7 134 29 ILL 72.5 40 
24 Jan 90 37 39 48.7 76 44 WS) 69.8 63 
31 Jan 90 42 23 64.6 65 31 4 88.6 35 

3 Feb90 39 46 45.9 85 35 10 77.8 45 
10 Feb 90 37 28 56.9 65 61 18 77.2 7) 


SOOrFrFNKFNONAANO 
_ 
PS 
(op) 


14 Feb90 56 39 58.9 95 at 8 90.6 85 181 
21Feb90 74 40 64.9 114 89 26 77.4 115 231 
28 Feb 90 29 ll 72.5 40 18 3 85.7 2] 62 

7 Mar 90 17 i) 77.3 22 ts) 4 55.6 ) 32 
14 Mar 90 q 6 53.8 13 3 0 100 3 16 
21 Mar90 = 12 0 100 12 I 1 50.0 2 14 


Totals 443 319 58.1 762 408 110 78.8 518 
% 6 = 6/(6 + 2); X = 6 + 2 (u = untagged and t = tagged); ? = sex undetermined; T = Xy + Xt + ?. 


bo 
bo 
-— 
oo 
S 
1X2) 


= 1, P < 0.68) indicates that the birds were not selective based on the 
butterfly’s sex. 

The proportion (17.7%) of previously tagged butterflies (recaptured 
butterflies in Table 2) in the sample of butterflies being tagged (total 
captured in Table 2) was compared with the proportion (31.3%) of 
tagged butterflies found in the preyed upon butterflies collected (Table 
4). The data strongly suggest that the birds were selecting tagged but- 
terflies (Chi-square = 130.70, df = 1, P < 0.001). The data also were 
separated on a weekly basis and a paired Student’s t-test was calculated. 
The data indicate that there was a very high probability that the tags 
increased the chances of avian mortality at this site ({ = —3.86, n = 9, 
P = 0.0048). 


DISCUSSION 


I first noted predation at the colony late in the season in 1988-89. 
This and the fact that I looked for it regularly beginning on 13 Sep- 
tember 1989 and did not notice it until 3 January 1990 suggests that a 
learning process was involved. Rufous-sided towhees were seen on sev- 
eral occasions earlier in the season, although it is uncertain whether it 
was the same pair seen throughout the study. But if it was, they were 
not eating the monarchs. Once they learned to eat monarchs, the re- 
source was exploited as shown in Tables 3 and 4. 

Field observations indicate that monarchs are difficult to spot when 
roosting in trees. But as sunrise approaches, they begin to open their 
wings and shiver, or after sunrise, they open their wings to bask to 
warm up to flight temperature. It was at this time that the towhees 


VOLUME 48, NUMBER 2 153 


began to find and attack the butterflies in the trees. But once the 
monarchs became warm enough to fly, towhee predation quickly ceased. 
Rufous-sided towhees belong to a ground feeding and seed eating sub- 
family of birds (Emberizinae); thus, the lack of success in sallying is 
expected once the butterflies are capable of flying (Bent 1968). 

In Mexico, Brower and Calvert (1985) found that birds preyed se- 
lectively upon males either because they had a lower fat content, higher 
wet weight, and/or a lower cardenolide content. This was not the case 
in this study, as the birds ate males in the proportion that they were 
found in the colony. Certainly the fact that the towhees are cueing in 
on the tagged monarchs affects the data. Since there is a difference 
between California and Mexican sites relative to the proportion of 
monarchs that are emetic as well as in their emetic toxicity (Brower & 
Moffitt 1974, Fink & Brower 1981, Brower & Fink 1985), the same 
mechanisms working in Mexico may not apply in California. Fink et 
al. (1983) have data suggesting that rufous-sided towhees in Mexico are 
not as sensitive to cardenolides, so the lack of prey selectivity by sex is 
consistent. 

Brower and Calvert (1985) estimated a 9.04% annual morality of 
butterflies killed by birds in one overwintering Mexican colony. Bell 
and Dayton (1986) reported annual mortality due to chestnut-backed 
chickadees to range from 0.345-2.21% in California. This study con- 
servatively estimated a 6.51-7.43% annual mortality. 

Bell and Dayton (1986) and Brower and Calvert (1985) found that 
predation intensity was higher on colder days in California and Mexico, 
respectively. The lowest predation rate during this study was 14.6 but- 
terflies per day during 24-31 January. During this week, an onsite 
hygrothermograph showed that there were two 36 hour periods when 
the temperature never fell below 19°C, well above monarch flight tem- 
perature of 18°C. There were essentially five days of warm Santa Ana 
conditions in southern California. The high temperature kept the but- 
terflies constantly agitated making it difficult for the towhees to catch 
them. On one such morning, butterflies were able to fly at 0400 h when 
disturbed. 

Based on observations of Bell and Dayton (1986) and Calvert et al. 
(1979), predators leave a characteristic signature on monarch remains. 
Perches regularly used by the towhees often had the remains (i.e. wings) 
of five or six butterflies in the litter under them. Brower et al. (1985) 
found such caches to be associated with small mammal predators in 
Mexico. 

Orioles, grosbeaks, and chickadees feed in the canopy where the 
clusters are, so body parts and wings simply rain down onto the ground. 
Calvert et al. (1979) and Fink and Brower (1981) described the different 


154 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


methods used by orioles and grosbeaks that feed on monarchs. Bell and 
Dayton (1986) found the chickadees’ mode of feeding is similar to that 
of orioles. As a comparison, yellowjacket predation involves biting off 
and carrying away the head and abdomen leaving the thorax with the 
four wings still attached (Leong et al. 1990, Sakai pers. obs.) 

The characteristic signature of rufous-sided towhees is a cluster of 
four wings found in the litter lacking a body. The heads occasionally 
are found but are difficult to find in the litter. I suspect that the towhee 
eats the entire body (head, thorax, and abdomen). Occasional heads in 
the litter are an artifact of the way the towhees kill the butterflies. 

Forty dead, intact butterflies were found in the litter during the 
course of the study. Towhees were observed attacking and picking up 
butterflies but later dropping and ignoring them. An untested possibility 
is that these were highly emetic individual butterflies. Mexican and 
California monarchs are different in their cardenolide biology, which 
is expected since they feed on different milkweed species. Brower and 
Moffitt (1974) report California monarchs were 4.6 to 6.5 times more 
emetic than their Mexican counterparts, and Brower and Fink (1985) 
report that 49% of California monarchs contain enough cardenolides 
to make them emetic to birds while only 10% of the Mexican monarchs 
were considered emetic. Brower and Calvert (1985) suggest that in 
Mexico birds feed cyclically because they accumulate cardenolides and 
must periodically desist to purge their bodies of these toxins. Based on 
these findings, one would expect less avian predation in California 
compared to Mexico, since California monarchs are both more emetic 
and have a greater percentage of emetic butterflies. 

Towhees may have learned to distinguish between emetic and non- 
emetic butterflies. Since the cardenolides are concentrated in the wings 
and exoskeleton, towhees may avoid eating, or learn not to eat, the most 
toxic ones as orioles do in Mexico. Assuming attacked but uneaten 
butterflies are the most toxic, the percent of emetic butterflies in this 
population is only 3.07% (40/1302). This is quite different from the 
findings of Fink and Brower (1981) and Brower and Fink (1985) of 
49% emetic butterflies in California. There are several possibilities. 
Towhees are able to distinguish the butterflies without killing the mon- 
archs. Towhees may be like grosbeaks and may be relatively insensitive 
to high doses of cardenolides. It also may be that the percentages of 
emetic butterflies and/or the degree of emetic strength found by Brower 
and his co-workers are not consistent throughout California. These 
questions certainly deserve investigation. , 

Brower and Calvert (1985) found that grosbeaks and orioles ate more 
males than females in the Mexican colonies, as females had 4% more 
lipid but a higher cardenolide concentration. Brower and Moffitt (1974) 


VOLUME 48, NUMBER 2 155 


found that in California males are heavier but females had a higher 
cardenolide concentration, meaning that it would be more advantageous 
to eat male butterflies. In this study, more males were eaten, but this 
may have reflected the skewed sex ratio in the colony rather than any 
selection of one sex over another. 

The actual impacts of butterfly tagging have never been addressed. 
This study indicates that tagged monarchs are at greater risk of pre- 
dation by rufous-sided towhees in the overwintering colony. It may be 
that the white tags serve as a flag to attract the birds and the birds 
learn to cue in on the tags. Tagged monarchs are certainly quite visible 
in the trees even to the naked eye. On the other hand, tagged wings 
are also much more visible in the litter. How much these tags serve to 
attract predators once the butterfly leaves the colony is unknown. Cer- 
tainly the migratory nature of the monarch would prevent predators 
from learning to cue in on the tags. 


(CONCLUSIONS 


A single pair of rufous-sided towhees was found to be an important 
predator on monarch butterflies at this southern California overwin- 
tering colony site, eating as many as 11.4 butterflies per bird per day, 
representing a 6.51-7.43% annual mortality. The predation rate seemed 
to be correlated with weather conditions with warmer weather resulting 
in lower predation rates. Towhees did not select prey by sex but were 
found to select a higher proportion of tagged monarchs. The charac- 
teristic pile of four wings may be used as a signature of rufous-sided 
towhees for determining them as a specific avian predator. 


ACKNOWLEDGMENTS 


I would like to thank J. Donahue, K. Garrett, L. Kiff, and K. Leong who commented 
on the early version of this paper. I am grateful for the helpful comments of W. Calvert 
and L. Brower as this manuscript evolved. R. Cobb provided proper terminology for 
avian behavior. I would like to thank the owner and caretaker of KSP for allowing me 
access to the site day or night. Partial funding for the field work was provided by Santa 
Monica College from California Assembly Bill 1725 (AB1725). Invaluable field assistance 
was provided by J. Ho and V. Sakai. 


LITERATURE CITED 


BELL, E. A. & J. J. DAYTON. 1986. Predation on monarch butterflies (Danaus plexippus ) 
by chestnut-backed chickadees (Parus rufescens) at a California overwintering site. 
In Donahue, J. P. (ed.), Abstracts on the Second International Conference on the 
Monarch Butterfly (MONCON-2). 2-5 September 1986. Natural History Museum of 
Los Angeles County, Los Angeles, California. 21 pp. 

BENT, A.C. 1968. Life history of North American birds. Cardinals, grosbeaks, buntings, 
towhees, finches, sparrows, and their allies; order Passeriformes, family Fringillidae. 
3 vol. Dover Publ., New York. 1889 pp. 


156 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


BROWER, L. P. & W. H. CALVERT. 1985. Foraging dynamics of bird predators on 
overwintering monarch butterflies in Mexico. Evolution 39:852-868. 

BROWER, L. P. & L. S. FINK. 1985. A natural toxic defense system: cardenolides in 
butterflies versus birds. Ann. N.Y. Acad. Sci. 443:171-188. 

BROWER, L.P., B. E. HORNER, M. M. Marry, C. M. MOFFITT & B. VILLA R. 1985. Mice 
(Peromyscus maniculatus labecula, P. spicelegus, and Microtus mexicanus salvus) 
as predators of overwintering monarch butterflies (Danaus plexippus) in Mexico. 
Biotropica 17:89-99. 

BROWER, L. P. & C. M. MorFitT. 1974. Palatability dynamics of cardenolides in the 
monarch butterfly. Nature 249:280-283. 

CALVERT, W. H., L. E. HEDRICK & L. P. BROWER. 1979. Mortality of the monarch 
butterfly (Danaus plexippus L.): Avian predation at five overwintering sites in Mexico. 
Science 204:847-851. 

FINK, L. S. & L. P. BROWER. 1981. Birds can overcome the cardenolide defence of 
monarch butterflies in Mexico. Nature 291:67-70. 

FINK, L. S., L. P. BROWER, R. WAIDE & P. R. SPITZER. 1983. Overwintering Monarch 
butterflies as food for insectivorous birds in Mexico. Biotropica 15:151-152. 

GLENDINNING, J. I., A. ALONSO M. & L. P. BROWER. 1988. Behavioral and ecological 
interactions of foraging mice (Peromyscus melanotis) with overwintering Monarch 
butterflies (Danaus plexippus) in Mexico. Oecologia 75:222-227. 

LEONG, K., D. FREY & C. NAGANO. 1990. Wasp predation on overwintering monarch 
butterflies (Lepidoptera: Danaidae) in Central California. Pan-Pacif. Entomol. 66: 
326-328. 

REMSEN, J. V. & S. K. ROBINSON. 1990. A classification scheme for foraging behavior 
of birds in terrestrial habitats. In Jehl, J. R., Jr. (ed.), Studies in Avian Biology 13: 
144-160. 

TusKEs, P. M. & L. P. BROWER. 1978. Overwintering ecology of the monarch butterfly, 
Danaus plexippus L., in California. Ecol. Entomol. 3:141-153. 

URQUHART, F. A. 1960. The monarch butterfly. University of Toronto Press, Toronto, 
Canada. 361 pp. 

. 1976. Found at last: The monarch’s winter home. Nat. Geogr. 150(August): 

160-1738. 

1987. The monarch butterfly: International traveler. Nelson Hall, Chicago. 

232 pp. 


Received for publication 8 February 1992; revised and accepted 31 July 1993. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 157-165 


BEHAVIORAL RESPONSE OF MONARCH BUTTERFLIES 
(NYMPHALIDAE) TO DISTURBANCES IN THEIR 
HABITAT—A GROUP STARTLE RESPONSE? 


WILLIAM H. CALVERT 
Department of Zoology, University of Florida, Gainesville, Florida 32611 USA 


ABSTRACT. Monarch butterflies (Danaus plexippus L., Nymphalidae) overwinter- 
ing in fir forests of Mexico’s Transvolcanic Belt fall from their perches at once en masse 
in response to “disturbance” by people or other animals that traverse their colonies. 
Roosting butterflies were stimulated by movement, noise, wind and breath. Their response, 
as measured by the number falling from perches, indicates that only breath is over- 
whelmingly effective in initiating the massive roost disintegrations observed. The response 
is temperature dependent; it begins about 7°C and increases as temperature rises. This 
group behavior is likely an adaptation to confuse and disorient bird and possibly mouse 
predators, increase their reaction time thus allowing the escape of the prey. Advantage 
may accrue to an individual belonging to a group whose size enhances the startle-effect 
of roost disintegration. 


Additional key words: Danaus plexippus, Mexico, overwintering biology, predation, 
protean behavior. 


Monarch butterflies (Danaus plexippus L., Nymphalidae) overwinter 
in huge aggregations numbering in the millions in the high altitude fir 
forests of Mexico’s Transvolcanic Belt (Calvert & Brower 1986). When 
people or other animals pass through the colonies, butterflies may drop 
from their clusters en masse sometimes in great numbers. Their col- 
lective falling was named “cascading” by K. Brugger, one of the Mex- 
ican monarch colony discoverers (Urquhart 1976). The behavior is not 
a passive event (Brower & Calvert 1985). The butterflies actively use 
their wings to cast themselves off their perches in a process that during 
warm weather sometimes appears explosive. The process may involve 
a few butterflies to hundreds of thousands of butterflies. When the 
wings are opened in the process of pushing off, the bright orange dorsal 
surface is suddenly exposed to view. The effect of tens or even hundreds 
of thousands of butterflies exposing these surfaces at once and falling 
from their perches is startling and beautiful. 

Guards and posted signs at two overwintering colonies open to tourists 
located within the ejidos (collective farms)—Rosario, near Ocampo, 
Michoacan, and Macheros, near Donato Guerra in the state of Mexico— 
admonish the public not to make loud noises while in the colonies. This 
request is in part an attempt to preserve the spirit of sanctuary within 
the butterfly colonies. But its main purpose is derived from a commonly 
held belief among local people that the cascading behavior of the 
monarch butterfly is a response to noise, such as speech, made by the 
visitors as they pass through the colony. 

Although there is a strong association between the presence of people 


158 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


and cascading of butterfly clusters, the precise cause(s) of roost disin- 
tegration have not been investigated. In this paper I explore several 
likely causes. 


METHODS 


Ten trunk and ten bough clusters ranging in height from 1-2 m and 
containing several hundred butterflies were artificially “disturbed” by 
movement, noise, air currents, and human breath. The degree of dis- 
turbance was assayed by estimating the number of butterflies cascading 
out of the roost. The response of roosting butterflies to movement was 
measured by moving the 2.5 cm diameter x 20 cm long grey plastic 
casing of an Omega RH-21C temperature/humidity probe slowly past 
the clusters approximately 2 cm from the butterflies. The response to 
loud noise was measured by clapping the hands twice 5-10 cm from 
the cluster. The hands were clapped together so that air was directed 
towards the researcher, not towards the butterflies. To test for air move- 
ment, the research notebook was waved once back and forth past the 
cluster rapidly enough so that the wind generated caused a passive 
movement of the wings. These tests put the researcher and his assistant 
in close proximity to the butterfly clusters. To avoid the possibility of 
confounding the tests above with breath, we held our breath and pe- 
riodically exhaled away from the butterfly cluster. Disturbance from 
breath was created by the researcher breathing directly on a cluster 
from a distance of 20 cm. The four types of disturbance were always 
tested in the order described on each of the ten bough and trunk clusters. 
The experimentally induced cascading of butterflies had no effect on 
adjacent clusters as is sometimes observed during warm weather. 

The experiment was performed on 29 and 80 January 1988 at the 
Palomas overwintering colony located on the west face of the Cerro de 
las Palomas approximately 35 km southwest of Toluca in the state of 
Mexico (Calvert & Brower 1986). Temperature and humidity of am- 
bient air were recorded by an Omega Engineering, Inc. RH-21C tem- 
perature/humidity probe. 


RESULTS 
Disintegration of Roosts or Cascading Behavior 


In response to a stimulus, clustered butterflies (Fig. 1) cast themselves 
off their perches in near synchrony (Fig. 2). This behavior involves 
more than the opening of the tarsal claws that grip the substrate. Using 
their wings, they actively push against each other, tree trunks or sur- 
rounding foliage. After casting off, they fall en masse tumbling over 
one another, and over foliage and branches until they either gain control 


VOLUME 48, NUMBER 2 159 


RP 


Fics. 1, 2. 1, Monarch butterflies clustered on a bough of the oyamel fir (Abies 
religiosa). The tightly packed butterflies are oriented in an orderly manner with their 
wings upright. Original Kodachrome slide by W. H. Calvert. 2, A monarch roost in the 
process of disintegration. The butterflies are falling in random disarray from a position 
marked by the white arrow. Original Kodachrome slide by Carlos Gottfried. 


and fly or until they strike the ground. The phenomenon gives the 
impression of a stream of large colored particles pouring downward 
over branches and foliage (Fig. 2). Cascading in one area of the colony 
may trigger similar disintegration of roosts in other areas resulting in 
tens or hundreds of thousands of butterflies leaving their clusters in a 
half hour period. 

An extreme example of cascading behavior occurred on a warm, 
sunny day in February of 1989 at the Palomas Colony (Calvert & Brower 
1986). Cascading began in one cluster at ca. 1400 h. Sequential cas- 
cading of clusters throughout the colony continued sporadically for the 
next hour. By 1500 h a section of forest approximately 20 x 30 m was 
covered in a living carpet of monarch butterflies ca. 8 cm deep. Excited 
butterflies were unable to fly but opened and closed their wings, thus 
pushing and moving against one another giving the appearance of a 
writhing mat of color. On grades this behavior caused them to flow en 
masse down slope where they piled up against trees or other obstacles 
such as fallen logs, brush piles or rocks. After several minutes of pushing 


160 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


themselves about with their wings, unless stimulated by more cascading 
butterflies falling upon them, they ceased moving their wings and began 
to crawl. When they encountered vegetation, they crawled upward 
(Alonso-Mejia et al. 1992). 

The majority of cascading events are much less spectacular. Bird 
predators such as blackheaded grosbeaks (Pheucticus melanocephalus) 
and blackbacked orioles (Icterus galbula abeillei) induce butterflies to 
cascade from positions where they are feeding. Tens to hundreds of 
butterflies have been observed to fall. More fell during warm ambient 
temperatures than cold. 

Cascading behavior occurs most often when air temperatures are 
near flight threshold, between 138-15°C (Masters et al. 1988, Kammer 
1970) or higher. At high air temperatures (20—25°C), clusters sometimes 
appear to break up spontaneously and the disintegration of one cluster 
nearly always triggers the disintegration of others. At air temperatures 
below flight threshold, multiple disintegrations are rare. Butterflies from 
a cascading cluster spill onto the ground and flap their wings helplessly. 
If it is not too late in the day, most of them will crawl up onto foliage 
(Brower et al. 1977). If air temperatures are above flight threshold or 
if the butterflies have had an opportunity to bask and have raised their 
thoracic temperatures to flight threshold (Masters et al. 1988), the falling 
butterflies will fly before they reach the ground. Sometimes they fly 
off in the same direction making a striking formation in the air. 


Butterfly Response to Movement, Noise and Wind 


The response of clustered butterflies to ‘disturbances’ by movement, 
noise and wind were measured by counting the number cascading from 
the roost. There was very little response to the movement of the probe 
near the clustered butterflies (Table 1). Only on one occasion did a few 
butterflies fall from a cluster in response to probe movement. This 
occurred at the relatively high temperature of 9.8°C. There was even 
less response to a sharp clapping noise. Only one butterfly fell from a 
cluster (temperature = 4.9°C). More response was evident to the winds 
generated by the research notebook. Some wing opening occurred at 
low temperatures of 0.9°C. Movement involving wing opening and small 
shifts in position occurred at 8.4°C. A few butterflies fell from the roosts 
at 18°C and higher. 


Butterfly Response to Breath 


The major cause of clustered butterflies falling from their perches 
was breath (Table 1). Butterflies directly in the path of the breath 
stream cascaded from their perches. A regression of temperature against 


VOLUME 48, NUMBER 2 161 


TABLE]. Reaction of clustered overwintering monarch butterflies to movement, noise, 
air currents and breath. 


Type of disturbance 


Cluster Temp. |b 10s Wp ei Ap OO a Ae SS AN eS sy ine tee 

Time Exposure type (°C) (%) Motion Sound Wind Breath 
08:02 Shade Bough —0.8 61.1 nc nc nc Ur 
08:06 Shade Bough 0.1 63.2 ne nc nc 5+ 
08:09 Shade Bough 0.1 62.3 ne ne nc 4+ 
08:18 Shade Trunk 0.9 56.4 ne ne 1+ 2+ 
08:22 Shade Trunk 0.9 50.1 ne ne l+ 3+ 
10:27 Shade Trunk 4.9 41.1 ne Il ne 10 
10:31 Shade Trunk 4.9 43.4 ne ne nc WD) 
10:34 Shade Bough 5.6 42.0 ne nc nc 2 
10:38 Sun Trunk 155) B7/ Il ne nc ne 30-40 
10:44 Sun Bough 11.8 34.2 ne ne nc 10 
13:39 Shade Trunk 8.9 43.1 nc ne ne 40-50 
13:43 Shade Trunk 9.0 42.6 nc nc nc 40-50 
13:48 Dappled Trunk 9.3 41.5 8 ne nc 25-30 
13:54 Shade Trunk 8.3 45.8 ne ne ne 15-20 
18:58 Shade Trunk 8.4 43.8 ne nce movt 15-20 
14:02 Shade Bough 8.4 43.2 nc ne movt 40-50 
14:06 Shade Bough 9.0 43.2 ne ne ne ~100 
14:14 Sun Bough 13.0 30.4 ne nc 4 ~100 
14:18 Sun Bough 15.0 32.2 ne nc 8 200-300 
14:22 Sun Bough 15.0 30.3 ne ne nc 200-300 


ne = no change; mov t = movement without opening wings; + = opened wings but did not drop. 


the number of butterflies cascading showed the response to be tem- 
perature dependent (F = 38.8; P = 0.0001; r? = 0.82). At all but the 
lowest temperatures, tens to hundreds of butterflies cascaded from their 
clusters in response to breath. Extremely cold butterflies (air temper- 
ature is 1°C or lower) responded to breath by opening their wings 
without falling from their perch. A few fell from their perches between 
4.9°C and 5.6°C. At temperatures between 7.5°C and 9°C, 15 to 100 
butterflies fell from their perches. At temperatures greater than 9°C, 
numbers near a hundred or above fell. 

No differences were apparent between cluster types. Bough clustered 
butterflies were as likely to cascade from their perches as were trunk 
clustered butterflies. No butterflies left their perches when relative 
humidity was above 50%, but this is likely an artifact of the inverse 
relation between temperature and relative humidity (r = —0.92) with 
temperature being the primary determinant of the butterfly response. 


DISCUSSION 
These data indicate that neither noise nor movement in the vicinity 
of clusters caused the cluster disintegration or cascading of butterflies 
observed during visits to the monarch butterfly colonies. Air currents 
directed onto clustered butterflies also did not evoke much response. 


162 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Some component of breath other than air current appears to be the 
cause of the massive cascading response. The failure of the butterflies 
to respond to the close presence of the researcher near the cluster until 
breathed upon suggests that heat per se is not the cause of roost dis- 
integration. | 


Erratic Escape Behavior 


Cascading behavior of monarchs is clearly more than a simple escape 
behavior. It involves a rapid, at times explosive, expulsion of numerous 
animals from a roost nearly simultaneously. The cryptic coloration of 
the roost suddenly erupts into the vivid orange colors of collective dorsal 
butterfly surfaces. 

Cascading behavior is analogous to behaviors in other animals. Prey 
animals in many different taxonomic groups behave erratically when 
attacked by predators (Tinbergen 1951, Roeder 1962, Cott 1940, Mar- 
shall & Orr 1955). Such erratic escape behavior, called protean behavior 
(Chance & Russell 1959), functions as an antipredator device by con- 
fusing and disorienting the predator, increasing its reaction time and 
enhancing the survival of the prey (Humphries & Driver 1967). The 
erratic and unpredictable nature of the response makes learned coun- 
termeasures by the predator less likely (Humphries & Driver 1970). 

Cascading behavior is perhaps most analogous to behavior exhibited 
by certain marine organisms. Antarctic krill maintain synchronized 
swimming patterns in apparent response to rheotactic cues supplied by 
the wake of preceding animals (Hamner et al. 1983). When frightened, 
their synchronized swimming pattern breaks apart, and they disperse 
in random directions. Monarchs roost in a very orderly manner with 
their folded wings oriented vertically on boughs or tree trunks (Fig. 1). 
When disturbed, this oriented assemblage breaks apart; initially the 
once clustered group tumbles towards the ground (Fig. 2), and if warm 
enough, flies off in random disarray. If they are able to fly, some sort 
of orderly flight pattern, likely towards a bright sky, is rapidly obtained. 

The elicitation of antipredator responses by mammalian breath may 
be of general occurrence in some groups of arthropods (Conner et al. 
1985). Certain millipeds react to breath by coiling, and they react to a 
warm surface by emitting defensive secretions. The tenebrionid beetle, 
Bolitotherus cornutus, everts quinone-producing glands when breathed 
upon. Some component of breath other than CO, seems to have elicited 
the response in the beetle. Possible components eliciting the response 
in monarchs include heat in the breath or one of the gaseous components 
of breath such as CO, or water vapor. Additional research is needed to 
determine which breath component evokes this startlingly beautiful 
defensive behavior in monarchs. 


VOLUME 48, NUMBER 2 : 163 


Microclimate and Bird Predation 


Because of their unique position in high altitude tropical forests and 
presence during the middle of the dry season, the microclimatic regimes 
of the overwintering colonies are precisely determined by the amount 
of radiation falling on the area. From the moment solar radiation strikes 
the area in the morning, parts of the forest begin to heat up and 
butterflies in the solar path or in the path of diffuse radiation are warmed 
to flight threshold and empowered to fly. Once solar input ceases in 
late afternoon, heat accumulated during the day rapidly radiates into 
the clear, dry sky. Temperatures drop precipitously and the butterflies 
rapidly lose the ability to respond to stimuli and to escape predators. 
If temperatures are cold so that the butterflies cannot fly and are forced 
to remain on the ground or on low foliage until the following day, they 
are subject to a possibly lethal combination of colder temperatures and 
higher humidities near the ground (Calvert & Brower 1981, Calvert & 
Cohen 1983) and to nocturnal mouse predation (Glendinning et al. 
1988). If temperatures are warm, or if it is early in the day so that a 
sun-fleck is likely to strike them, most butterflies will return to their 
elevated roosts before harm comes to them. However, even when no 
direct harm comes to them, they must use scarce lipid reserves to crawl 
up onto foliage or to fly back to their roosts (Masters et al. 1988). Because 
nectar sources are limited during the overwintering season (Brower & 
Malcolm 1991), this use of energy reserves could result in premature 
starvation. 

In Mexico’s overwintering sites bird, mainly blackheaded grosbeaks 
(Pheucticus melanocephalus) and blackbacked orioles (Icterus galbula 
abeillei), and mouse, mainly Peromyscus melanotis, predators killed 
an estimated 926,000 butterflies/ha in a 185 day season (Brower & 
Calvert 1985, Glendinning et al. 1988) which may have amounted to 
as much as 10% of the colony population (Calvert et al. 1988). Birds 
feed during the coldest times of the day usually from dawn to 0900 or 
1000 h and from ca. 16380 h till dark (Brower & Calvert 1985). Feeding 
early and late, birds are likely to encounter cold butterflies unable to 
move quickly to escape. Birds feeding when solar radiation is striking 
the colony may encounter butterflies with thoracic temperatures above 
ambient that are capable of vigorous movement and flight. Attacking 
while the butterflies are inactivated by the cold would be likely to 
insure minimum handling cost per effort (Krebs 1978, Brower & Calvert 
1985). If the majority of predators are birds which exhale upon clustered 
monarchs in the process of capturing them, a group startle response 
such as the cascading behavior designed to confuse and disorient the 
predator to increase its reaction time would be a beneficial adaptive 
response to the butterflies. 


164 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


These data suggest that the butterflies benefiting the most from the 
cascading response are those whose thoracic temperatures are between 
ca. 7°C and flight threshold (Table 1). At temperatures lower than this, 
it is more difficult for them to cast off their perches. At higher tem- 
peratures they can fly away to escape predation. Except for the coldest 
hours of early morning or late evening, most clustered butterflies are 
likely to experience temperatures in this range. (Exceptions include 
parts of November and March when ambient temperatures in the forest 
climb above flight threshold and cold overcast days of winter when 
ambient temperature never reaches 7°C.) Cascading behavior is unlikely 
to benefit butterflies being preyed upon by mice since nighttime tem- 
peratures when mice feed are nearly always colder than 7°C. The 
cascading response is most likely to benefit butterflies that are able to 
cast off of their perches en masse, but are unable to fly due to cold air 
temperatures. 

There remains the possibility that two different kinds of cascading 
phenomena occur in the monarch overwintering colonies. The dramatic 
disintegration of monarch roosting clusters involving tens of thousands 
of individuals may be an artifact of gregarious roosting and a different 
phenomenon from the smaller scale cascading response elicited by avian 
predators. These multiple cluster disintegrations are especially apparent 
during warm periods late in the season (February or March) and may 
be part of the colony breakup in preparation for the return migration 
to the north. Drawn to nectar and water sources at lower elevations, 
the colonies rapidly move down slope, often dividing into two or more 
segments (Calvert & Brower 1986). Mating activity is intensified and 
each day a portion of the colony leaves to begin its remigration north- 
ward (Van Hook 1998). 


ACKNOWLEDGMENTS 


I thank Alicia Nelson and David Richie for helping to collect the data. The ideas that 
led to these experiments were bantered around in campfire discussions near the Sierra 
Chincua research colony in Mexico. Some of the participants were Tonya van Hook, John 
Glendinning, Jim Anderson, Alfonso Alonzo, Alfredo Arellano and Lincoln Brower. John 
Alcock, Philip Russell, Steve Malcolm, Alicia Nelson, Lincoln Brower, Walt Sakai, and 
an anonymous reviewer made suggestions to improve the manuscript. This research was 
supported in part by Monarcha A.C. with funds from the World Wildlife Fund US and 
the Cecil B. Day Foundation of Atlanta, Georgia. 


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J. R. & N. B. Davies (eds.), Behavioral ecology: An evolutionary approach. Sinauer 
Associates, Sunderland, Massachusetts. 

MARSHALL, S. M. & A. P. OrR. 1955. The biology of the marine copepod Calanus 
finmarchicus (Gunnerus). Oliver and Boyd, Edinburgh. 188 pp. 

MasTERS, A. R., S. B. MALCOLM & L. P. BROWER. 1988. Monarch butterfly (Danaus 
plexippus) thermoregulatory behavior and adaptations for overwintering in Mexico. 
Ecology 69:458—467. 

ROEDER, K. D. 1962. The behaviour of free flying moths in the presence of artificial 
ultrasonic pulses. Animal Behaviour 10:300-304. 

TINBERGEN, N. 1951. The study of instinct. Oxford University Press, Oxford, England. 
228 pp. 

URQUHART, F. A. 1976. The overwintering site of the eastern population of the monarch 
butterfly (Danaus plexippus, Danaidae) in southern Mexico. J. Lepid. Soc. 30:153- 
158. 

VAN Hook, T. 1993. Non-random mating in monarch butterflies overwintering in 
Mexico, pp. 49-60. In Malcolm & Zalucki (eds.), Biology and conservation of the 
monarch butterfly. Natural History Museum of Los Angeles County, Contributions 
in Science, Los Angeles. 419 pp. 


Received for publication 6 June 1998; revised and accepted 18 September 1993. 


GENERAL NOTES 


Journal of the Lepidopterists’ Society 
48(2), 1994, 166-168 


THE CASE OF THE MISSING H: HELICONIUS CHARITHONIA 
(L., 1767), NOT “HELICONIUS CHARITONIA (L., 1767)” 


Additional key words: Nymphalidae, Heliconiini, nomenclature, taxonomy, spelling. 


Systematic nomenclature is perhaps the only truly typological part of our science, since 
it depends on published historical records and not on the vagaries of organismal variation. 
There are correct names, and there are incorrect names. This little note is about spelling, 
and thus may seem trivial and pedantic, but there are four reasons why my topic is 
noteworthy (if you will). First, the taxon in question is the type species of the celebrated 
genus Heliconius, which has enjoyed paradigmatic status in ecological genetics, historical 
biogeography, and community ecology for several decades. Second, the incorrect spelling 
of the name is in wide use: a quick scan through literature from the last ten years yields 
more than a dozen uses of charitonia and no uses of charithonia. This raises point three: 
as electronic reference data bases become more prevalent, alternate spellings of names 
must be eliminated, to promote efficient searching. Lastly, the story behind the confusion 
is entertaining, if nomenclatorially and historically convoluted. 

Papilio Heliconius charithonia was described as a species by Linnaeus in the 12th 
edition of Systema Naturae (1767), but, because he considered all butterflies to belong to 
the genus Papilio, the infrageneric name, Heliconius is invalid. (Interestingly, if Hel- 
iconius L. were valid, the type species would be H. ricini L., 1758 and not H. charithonia, 
L. 1767, but that’s another story.) For almost 150 years, authors attributed the name 
Heliconius to Latreille (either 1804 or 1805), with a variety of type species including 
charitonia (Fabricius, 1775)). 

In 1938, the great historian of lepidopteran systematic literature, Francis Hemming, 
recognized that Heliconius (Latreille, 1804) lacked a valid type species, and selected 
charitonia Fabr., which he equated with charithonia L. (Hemming 1933a). Immediately 
after publication of this designation, Hemming apparently discovered a prior binomial 
usage of Heliconius by Kluk (1802). He quickly published a new designation, this time 
listing charitonia L. as the type species, without the “h” (Hemming 1933b). By 1934, 
however, Hemming had returned to charithonia, and clearly acknowledged the differ- 
ences in spelling between the early authors, indicating his preference for the Linnaean 
use of the “h.”’ This decision is reiterated in his posthumous magnum opus (Hemming 
1967). 

Apparently unaware of Hemming’s efforts, Comstock and Brown specifically addressed 
the problem of the “h” again in 1950. They pointed out that the index of the 12th edition 
of Systema Naturae (1767) makes reference to charitonia, as does the 18th.edition (1790). 
They argue further that the name is etymologically derived from charites, Latinized 
from the Greek name for the Graces, and thus logically not containing the “h.” They 
claimed that since the two spellings are of equal age, and since no other revisers had 
addressed the issue, they could choose charitonia as the proper spelling. Amazingly, in 
their next paragraph, they coined yet another version of the name, charitonius, to produce 
gender agreement between the genus and the species. This paper appears to be the source 
of current ubiquitous usage of charitoni- (with the -a or -us suffix variously applied). 

Five years after Comstock and Brown tried to lay it to rest, the “h” issue rose up yet 
again in a case submitted to the International Commission on Zoological Nomenclature 
regarding the priority of names for a crustacean (Holthuis & Hemming 1956). It seems 
that Sicyonia thamar, the name given by Hiibner (1816) to what we recognize today as 
Heliconius sara (Fabr. 1793), had priority over a generic name used for a well known 
genus of prawn (Sicyonia, Milne Edwards 1830). Under its plenary powers, the Com- 
mission decided to sink Sicyonia Hiibner, in deference to the request to retain the popular 
usage of the name for the prawn, and because the butterfly name was not in use, the 


VOLUME 48, NUMBER 2 167 


species described by Hiibner being considered to belong to Heliconius. In an addendum 
to this decision, Hemming, the Secretary of the Commission, officially rejected Heliconius 
Latreille, 1804; Apostraphia Hubner, 1816; Heliconia Godart, 1819; and Heliconius L., 
1758. He also placed Heliconius Kluk, 1802 on the Official List of Generic Names in 
Zoology, and, crucial to the point of this note, he put charithonia L., 1767 on the Official 
List of Specific Names in Zoology (Melville & Smith 1987). 

Decisions made under the plenary powers of the I.C.Z.N. overrule previous arguments 
about nomenclature, and thus clearly invalidate Comstock and Brown’s (1950) arguments, 
whether they bear merit or not. It is arguable, however, that Comstock and Brown’s views 
are based on poor interpretations of the International Code of Zoological Nomenclature 
(Ride et al. 1985). Even if their dubious claim to First Reviser status is valid, giving them 
the prerogative to choose between alternate spellings in the original description (Article 
24(C)), the Code recommends that the spelling that appears first be chosen when it is not 
obviously wrong or will not fail to serve universality of nomenclature (Recommendation 
24(A)). Prior to Comstock and Brown's dictum, most major Heliconius systematists (Kirby 
1871, Riffarth 1901, Stichel 1906, Eltringham 1916, Seitz 1924, Neustetter 1929, and of 
course, Hemming 1938a, 1933b, 1934) used charithonia. Furthermore, it seems more 
appropriate to choose the name accompanying the original description than the name 
listed subsequently in an index, which might have been less subject to editorial scrutiny. 
Linnaeus personal copy of the 12th edition of Systema Naturae (1767), although filled 
with hand-written corrections and amendments, shows no suggestion that Linnaeus viewed 
charithonia as a misprint in his text. 

With regards to the etymology of the name, there is no extrinsic evidence to suggest 
that charitonia is derived from Charites. As Turner (1967) pointed out, many 18th century 
names do not derive unambiguously from Latin or Greek roots. Turner also chided authors 
for inappropriate masculinization of specific names to bring them into gender agreement 
with generic names. Article 31(B(i)) of the Code (Ride et al. 1985) explicitly states that 
names stand as first published, regardless of gender, unless the author specifically stated 
that the species name is an adjective modifying the generic name, which Linnaeus did 
not. 

So there we have it. Heliconius charithonia, described by Linnaeus (1767), designated 
as type species of the genus by the rightful First Reviser, Hemming (1933b), and placed 
on the I.C.Z.N. official list of generic and specific names by Holthuis and Hemming 
(1956). Yet every major guide to butterflies published since then has followed Comstock 
and Brown (1950), listing the species as ““charitonia,” (or even “charitonius,” employing 
their demonstrably incorrect masculinization). I hope that this minor, yet irritating detail 
will be corrected in future publications. 


LITERATURE CITED 


Comstock, W. P. & F. M. BRowN. 1950. Geographical variation and subspeciation in 
Heliconius charitonius Linnaeus (Lepidoptera, Nymphalidae). Am. Mus. Novit. 1467: 
1-21. 

ELTRINGHAM, H. 1916. On specific and mimetic relationships in the genus Heliconius, 
L. Trans. Entomol. Soc. Lond. 1916:101-148. 

FABRICIUS, J. C. 1775. Systema Entomologiae. Pp. 458-465. 

1793. Entomol. Syst. 3(1):167. 

GopakT, J. B. 1819. Ency. Méth. 9(1) (Ins.):203. 

HEMMING, F. 1933a. On the types of certain butterfly genera. Entomologist 66:196- 
200. 


1933b. Additional notes on the types of certain butterfly genera. Entomologist 

66:222-225. 

1934. The generic names of the Holarctic butterflies I. 1758-1863. British 

Museum (Natural History). 184 pp. 

1967. The generic names of the butterflies and their type-species (Lepidoptera: 
Rhopalocera). Bull. Br. Mus. Nat. Hist. (Ent.), Suppl. 9, 509 pp. 

Ho.tuHuis, L. B. & F. HEMMING. 1956. Opinion 382: Validation under the Plenary 
Powers of the generic name “Sicyonia”’ Milne Edwards (H.) 1830 (class Crustacea, 


168 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


order Decapoda) and action consequential thereon, pp. 43-58. In Hemming, F. (ed.), 
Opinions and declarations rendered by the International Commission on Zoological 
Nomenclature 12(3). 

HUBNER, J. 1816. Verz. Bekannt. Schmett. 1:13. 

KirBy, W. F. 1871. A synonymic catalogue of diurnal Lepidoptera. John Van Voorst, 
London. 690 pp. 

KLuK, K. 1802. Zwierz. Hist. Nat. Pocz. Gospod. 4:82. 

LATREILLE, P. A. 1804. Nouvelle Dictionaire d’Histoire Naturelle 24 (Tab.):185, 199. 

1805. Histoire Naturelle Genérale et Particulaire, des Crustacés et des Insectes. 
14:108. 

LINNAEUS, C. 1758. Systema Naturae. 10th ed. 464. 

1767. Systema Naturae. 12th ed. 1(2):757. (Microfiche facsimile of Linnaeus’ 
personal copy. London. British Museum (Natural History) and the Linnean Society, 
1991). 

MELVILLE, R. V. & J. D. D. SMITH (eds.). 1987. Official lists and indexes of names and 
works in zoology. International Trust for Zoological Nomenclature and the British 
Museum (Natural History), London. 209. 

MILNE EDWARDS, H. 1830. Ann. Sci. Nat., Paris (1) 19:339. 

NEUSTETTER, H. 1929. Nymphalididae: subfam. Heliconiinae. Lepidopterorum Cata- 
logus. 36:1-136. 

RIDE, W. D. L., C. W. SABROSKY, G. BERNARDI & R. V. MELVILLE (eds.). 1985. Inter- 
national code of zoological nomenclature. International Trust for Zoological Nomen- 
clature and the British Museum (Natural History), London. 338 pp. 

RIFFARTH, H. 1901. Die Gattung Heliconius Latr.: Neu bearbeitet und Beschreibung 
neuer Formen. Berl. Entomol. Zeit. 46:25-183. 

SEITZ, A. 1924. Subfamily: Heliconiinae. Macrolepidoptera of the world: The American 
Rhopalocera. Alfred Kernen, Stuttgart 5:593-597. 

STICHEL, H. 1906. Lepidoptera Rhopalocera fam. Nymphalidae subfam. Heliconiinae. 
Genera Insectorum 37:1-74 (6 plates). 

TURNER, J. R. G. 1967. Goddess changes sex, or the gender game. Syst. Zool. 16:349- 
300. 


ANDREW V. Z. BROWER, Section of Ecology and Systematics, Cornell University, 
Ithaca, New York 14853-2701, USA (current address: Department of Entomology, Amer- 
ican Museum of Natural History, Central Park West at 79th St., New York, New York 
10024-5192). 


Received for publication 15 July 1993; revised and accepted 30 October 1993. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 168-170 


NATIVE PIERINE BUTTERFLY (PIERIDAE) ADAPTING TO 
NATURALIZED CRUCIFER? 


Additional key words: Brassicaceae, diet breadth. 


Native butterflies encounter naturalized plants related to their hosts as one consequence 
of Palearctic weeds spreading throughout North America. Sometimes these plants are 
incorporated into the butterfly diet and permit a longer flight season (e.g., Pieris napi 
microstriata on watercress: Shapiro 1975; Papilio zelicaon on sweet fennel: Sims 1983, 
Tong & Shapiro 1989, Shapiro in press). In other cases, females do not lay eggs on the 
naturalized plant so that it is not used, even though it can support complete larval 
development (e.g., Colias philodice and crown vetch: Karowe 1990). A third alternative 


VOLUME 48, NUMBER 2 169 


is that females lay eggs on naturalized plants, but larvae die before completing devel- 
opment (e.g., Pieris napi oleracea, P. n. marginata and P. virginiensis on garlic mustard: 
Bowden 1971; P. n. macdunnoughii and P. occidentalis on pennycress: Chew 1977; P. 
napi oleracea on wintercress: Chew 1981). 

Where naturalized plants are attractive to ovipositing females but larvae fail to develop 
successfully on them, the plants can function as a population sink so that eggs laid on 
these plants are lost from the population. If appropriate genetic variation exists in the 
population, selection favors both females that discriminate against the unsuitable plant, 
and feeding larvae that are able to develop on it successfully (Chew 1977). We report 
here on a population of P. napi oleracea Harris that may be adapting to a biennial weed 
of Palearctic origin, garlic mustard, Alliaria petiolata (Bieb.) Cavara & Grande (Brassi- 
caceae) (nomenclature follows Gleason & Cronquist 1991). 

Pieris virginiensis and P. napi oleracea fly together in beech-maple-hemlock woods 
near Lee and Dalton (Berkshire Co., Massachusetts) and Sandgate (Sandgate Co., Ver- 
mont). At Lee, Roger W. Pease, Jr. (RWP) previously observed both species ovipositing 
on garlic mustard and found eggs and small larvae of P. n. oleracea on this plant in July 
(RWP in litt. and pers. comm.). Rearing these eggs on garlic mustard produced adults 
with summer brood P. n. oleracea phenotypes (RWP pers. comm.). Garlic mustard stands 
are dense and extensive at this locality and large stands of toothwort, Cardamine diphylla 
(Michx.) A. Wood also are present. Occasional plants of watercress, Rorippa nasturtium- 
aquaticum (L.) Hayek, and cuckoo-flower, Cardamine pratensis L., also occur. This site 
is 17 km from the Dalton site, where both butterflies fly together (Chew 1980) and utilize 
C. diphylla. At Dalton, on 19 August 1993, we observed garlic mustard only in two very 
small stands near a parking lot; one stand lacked rosettes, indicating that no seeds ger- 
minated on this spot during the past spring—a probable indication of recent establishment. 
To our knowledge, garlic mustard has not been reported previously at the Dalton site. 
At both sites, toothwort is present above-ground in the spring, but by mid-July its leaves 
are severely yellowed and unsuitable as Pieris food. By contrast, garlic mustard flowering 
stalks remain green in mid-July, and rosettes remain green throughout the summer. 

We confirmed RWP’s observations of P. virginiensis and P. n. oleracea females ovi- 
positing on garlic mustard in 1992 and 1993. On 7 May 1993 we collected a single female 
P. n. oleracea. We attempted to rear some of this female’s offspring on garlic mustard. 
Of 34 newly hatched first instar larvae reared on this plant, 14 pupated (8 female, 6 
male). Pupal weights for both sexes were not significantly different from weights for sibs 
reared on hare’s-ear mustard. However, larval developmental times (days from egg hatch 
to pupation) differed significantly for individuals reared on garlic mustard compared to 
sibs reared on hare’s-ear mustard (range on garlic mustard = 16-30 d compared to 14- 
17 d for hare’s-ear mustard; mean + SD for females on garlic mustard = 23.4 + 3.54 d 
compared to 15.5 + 1.64 d on hare’s-ear mustard; mean + SD for males on garlic mustard 
= 19.0 + 2.90 d compared to 14.3 + 0.58 d on hare’s-ear mustard). All 14 pupae emerged 
after 6-8 days. Whether these adults were fertile is not known, because they were then 
placed in a common cage with sibs reared on hare’s-ear mustard. The collective cage 
produced fertile eggs but we cannot be certain whether any came from individuals reared 
on garlic mustard. 

Garlic mustard is a widely used and highly suitable host of P. napi and many related 
species in Europe (Bowden 1971, A. Porter in litt.). But previous attempts to rear North 
American pierines on garlic mustard have failed, usually in the first or second instar 
(Bowden 1971; our unpubl. data). F, hybrids between North American and English 
pierines develop successfully on this plant (Bowden 1971). Our only previous success 
involved 5 P. n. oleracea (3 females, 2 males) derived from a combined breeding stock 
from Lee and central Vermont (near Hancock, Hancock Co., Vermont). These individuals 
took longer to develop from egg hatch to pupation, but their pupal weights were similar 
compared to other members of the colony reared on hare’s-ear mustard. 

Recent work on the behavior of ovipositing P. n. oleracea from central Vermont shows 
that these butterflies are more strongly stimulated by alkenyl glucosinolates such as allyl 
glucosinolate (sinigrin) than by some other glucosinolates (Chew & Renwick in press). 
Because allyl glucosinolate is prominent in the glucosinolate profile of garlic mustard 


170 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


leaves (Chew, F. S., unpubl. data; Renwick, J. A. A., unpubl. data), this plant’s attrac- 
tiveness to ovipositing P. n. oleracea is not surprising. Individuals that develop successfully 
on garlic mustard would be able to use extensive stands of this naturalized host in addition 
to their toothwort host, and the two hosts together presumably could support a larger 
butterfly population. Whether the observed variation in larval success on garlic mustard 
is a remnant of genetic heritage from a pierine ancestral to both North American and 
European P. napi lineages, or whether it is newly arisen in the Berkshire Co., Massachusetts 
region, is unknown. 

We thank R. W. Pease, Jr. for introducing us to P. virginiensis and P. n. oleracea near 
Lee. We thank R. W. Pease, A. M. Shapiro, and A. Porter for sharing their observations 
and helpful discussion with us. We thank NSF (IBN-91-08987) and the Arabis Fund for 
financial support. 


LITERATURE CITED 


BOWDEN, S. R. 1971. American white butterflies (Pieridae) and English food-plants. J. 
Lepid. Soc. 25:6-12. 

CHEW, F. S. 1977. Coevolution of pierid butterflies and their cruciferous food plants. 
II. The distribution of eggs on potential hostplants. Evolution 31:568-579. 

1980. Natural interspecific pairing between Pieris virginiensis and P. napi 

oleracea (Pieridae). J. Lepid. Soc. 34:259-260. 

1981. Coexistence and local extinction in two pierid butterflies. Am. Nat. 118: 
655-672. 

CHEW, F. S. & J. A. A. RENWICK. in press. Hostplant choice in Pieris butterflies. In 
Cardé, R. T. & W. J. Bell (eds.), Chemical ecology of insects II. Chapman & Hall, 
New York. 

GLEASON, H. A. & A. CRONQUIST. 1991. Manual of vascular plants of northeastern 
United States and adjacent Canada. 2nd ed. New York Bot. Gard., Bronx. 

KAROWE, D. N. 1990. Predicting host range evolution: Colonization of Coronilla varia 
by Colias philodice (Lepidoptera: Pieridae). Evolution 44:1637-1647. 

SHAPIRO, A. M. 1971. Occurrence of a latent polyphenism in P. virginiensis (Lepidop- 
tera: Pieridae). Entomol. News 82:13-16. 

1975. The role of watercress (Nasturtium officinale) as a host of native and 

introduced pierid butterflies in California. J. Res. Lepid. 14:158-168. 

in press. From the mountains to the prairies to the oceans white with foam: 

Papilio zelicaon makes itself at home. In Walker, R. B. & A. R. Kruckberg (eds.), 

Ecogeographical races: Turesson to the present. Am. Assoc. Adv. Sci., Washington, 


Sims, S. R. 1983. Inheritance of diapause induction and intensity in Papilio zelicaon. 
Heredity 51:495-500. 

TonG, M. L. & A. M. SHAPIRO. 1989. Genetic differentiation among California popu- 
lations of the anise Swallowtail butterfly, Papilio zelicaon lucas. J. Lepid. Soc. 48: 
217-228. 


A. V. COURANT, A. E. HOLBROOK, E. D. VAN DER REIJDEN, AND F. S. CHEW, De- 
partment of Biology, Tufts University, Medford, Massachusetts 02155, USA. 


Received for publication 4 September 1993; revised and accepted 1 January 1994. 


Journal of the Lepidopterists’ Society 
48(2), 1994, 171-172 


IMPLICATIONS OF INTRODUCED GARLIC MUSTARD 
(ALLIARIA PETIOLATA) IN THE HABITAT OF 
PIERIS VIRGINIENSIS (PIERIDAE) 


Additional key words: host expansion, conservation. 


Pieris virginiensis (W. H. Edwards) (Pieridae) is an uncommon and local univoltine 
member of the P. napi (L.) group from eastern North America, confined to forest un- 
derstory habitats where its primary hosts, Dentaria spp. (Brassicaceae), have relatively 
dense populations (Opler & Krisek 1984). Most other populations of the napi group are 
multivoltine, as are other Pieris species (brassicae [L.], rapae [L.], manni Mayer, etc.). 
The other univoltine napi-group members are confined to habitats with short growing 
seasons, and typically hybridize in ecotones with multivoltine races (Petersen 1963; my 
unpubl. data). Allozyme data confirm that virginiensis is very close to other North 
American napi-group taxa (Geiger & Shapiro 1992). Univoltinism in P. virginiensis is 
presumed to be a derived adaptation to early senescence of Dentaria because summer 
generations would not have adequate host resources to complete larval growth (Shapiro 
1971). 

Garlic mustard, Alliaria petiolata (M.B.) (Brassicaceae), is an aggressive forest-edge 
and understory plant that was introduced into northeastern North America from Europe, 
and has become widespread in the midwestern U.S. since around 1950; there is interest 
in control of its spread (Willis 1992). Alliaria germinates in the spring and summer and 
forms a large, overwintering rosette, which then bolts in the early spring and blooms 
from late March to July. Plants with edible foliage can thus be found throughout the 
year. 

At the Cedar Bluffs Nature Preserve in Monroe Co., Indiana, Dentaria laciniata Willd. 
is native and patchily distributed, whereas Alliaria petiolata is more widespread on the 
site. On 1 May 1998, Dentaria had already set seed, but the Alliaria were still in bloom. 
At approximately 1400 h, I observed a female P. virginiensis while she laid 6 eggs over 
a 10 minute period. Host-searching behavior was typical for Pieris: she flew slowly and 
frequently contacted leaves and flowers of a variety of plant species, but flew up im- 
mediately after contact with non-cruciferous leaves. When she contacted an Alliaria, 
sometimes by diving at the flower (suggesting visual cues were involved), she began 
searching and alighting on the lower leaves, ~1-2 dm above the ground. In most cases, 
she would lay a single egg on the underside of a leaf at this height, but several plants 
were rejected. After oviposition, she always flew at least several meters before searching 
for the next host. She did not contact any Dentaria during this time. After collecting this 
female as a voucher (retained in my personal collection), I searched Dentaria, and 
occasional Arabis (Brassicaceae) plants for 10 minutes, and did not find any eggs or larvae. 
The female subsequently laid eggs on Alliaria in captivity; these eggs were reared into 
the second instar with moderate mortality before fieldwork overseas terminated the study. 

There are three possible outcomes of the use of Alliaria as an oviposition site by P. 
virginiensis, and they depend on the palatability of the plant to the larvae. If the plant 
is toxic, then there are two possible outcomes: the virginiensis populations will decline 
because females waste eggs on Alliaria; and/or there will be selection on females to avoid 
Alliaria together. However, if Alliaria is palatable, even to some larvae in the population, 
then virginiensis will probably evolve a broader host range because Alliaria is a more 
widespread and reliable resource than Dentaria. 

Isolated populations of virginiensis are likely to be in danger of extinction if Alliaria 
is toxic to the larvae. Pieris virginiensis maintains relatively small and localized popu- 
lations, and does not often fly outside the forest understory. Alliaria is likely to be much 
more apparent than Dentaria in virginiensis habitat, both because Alliaria reaches high 
abundance over larger areas than Dentaria, and because Alliaria have larger plants with 
a longer flowering season. Female virginiensis searching for crucifers are likely to en- 
counter and oviposit on Alliaria more often than Dentaria, and perhaps be prone to fly 


72 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


out of Dentaria patches while continuing to discover Alliaria plants. The resulting decline 
is likely to lead to extinctions of already small virginiensis populations before the ability 
to discriminate can evolve. Even if the ability to discriminate does evolve locally, it is 
unlikely to spread rapidly and “rescue” other populations because of the philopatric 
tendencies of virginiensis. 

On the other hand, it would be a welcome development if P. virginiensis were to 
expand its host range to include Alliaria. It would give P. virginiensis a better foothold 
where it does live, and perhaps allow it to expand its range of suitable habitats. In Europe, 
Alliaria petiolata is a common host of Pieris napi (Petersen 1963, Bowden 1971; my 
unpubl. data), but in North America, P. napi oleracea Harris has reportedly been un- 
successful on Alliaria (Bowden 1971), although I have frequently found P. rapae to use 
it (unpubl. data). Incipient host-range expansion is also of interest to population and 
community ecologists: because Alliaria foliage can be found in abundance throughout 
the year, natural selection would favor the return of Pieris virginiensis to its ancestral, 
multivoltine life cycle. 

Population studies following the historical progress of the interactions between Alliaria 
and P. virginiensis, regardless of the outcome, would be quite relevant to our under- 
standing of how host selection and host use evolves in natural populations. For example, 
Rhagoletis pomonella (Diptera: Tephritidae) is presumed to have switched from hawthorn 
to apple in New York State, whereupon the apple-feeding race spread over the north- 
eastern U.S. (Bush 1968). Even though this case is among the best studied, the evidence 
for this scenario is weak because historical data are scant, and the switch may as easily 
have occurred more or less simultaneously in several parts of the species’ range. Courant 
et al. (1994) independently report interactions between Alliaria and P. napi oleracea and 
Alliaria and P. virginiensis in New England, with similar observations to mine from 
Indiana; notably, New England populations have been in the presence of Alliaria longer 
than midwestern populations and may have had an evolutionary “head start.” Strong 
selection by Alliaria is probably occurring over much of the range of virginiensis, and 
simultaneous observations of the interactions in geographically separated populations are 
needed to help understand processes involved in the evolutionary response to this selection. 

This note has benefitted from discussions with Francie Chew and Art Shapiro, and two 
anonymous reviewers comments. Francie was kind enough to send me her unpublished 
manuscript. Thanks are due to Mark Dybdahl and Lynda Delph for accommodations in 
Bloomington, and to Lynda for identifying the Dentaria. 


LITERATURE CITED 


BOWDEN, S. R. 1971. American white butterflies and English food-plants. J. Lepid. Soc. 
25:6-12. 

BusH, G. 1968. Sympatric host race formation and speciation in frugivorous flies of the 
genus Rhagoletis (Diptera, Tephritidae). Evolution 23:237-251. 

COURANT, A. V., A. E. HOLBROOK, E. D. VAN DER REIJDEN & F. S. CHEW. 1994. Native 
pierine butterfly (Pieridae) adapting to naturalized crucifer? J. Lepid. Soc. 48:168- 
7A), 

GEIGER, H. & A. M. SHAPIRO. 1992. Genetics, systematics and evolution of holarctic 
Pieris napi species group populations (Lepidoptera: Pieridae). Z. Zool. Syst. Evol.- 
forsch. 30:100-122. 

PETERSEN, B. 1963. Breakdown of differentiation between Pieris napi L. and Pieris 
bryoniae Osch. and its causes. Zool. Bidr. Uppsala 35:1-262. 

OPLER, P. A. & G. O. KRISEK. 1984. Butterflies east of the plains. Johns Hopkins Univ. 
Press, Baltimore. 

SHAPIRO, A. M. 1971. Occurrence of a latent polyphenism in Pieris virginiensis (Lep- 
idoptera: Pieridae). Entomol. News 82:13-16. 

WILLIS, J. 1992. Garlic mustard is hard to control. Trillium 1992:8-11. 


ADAM PorRTER, Department of Biological Sciences, Bowling Green State University, 
Bowling Green, Ohio 43403-0212, USA. 


Received for publication 8 May 1993; revised and accepted 20 November 1993. 
Date of Issue (Vol. 48, No. 2): 17 May 1994 


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196la. Some contributions to population genetics resulting from the study of 

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CONTENTS 


SAMPLING STRATEGIES FOR ESTIMATING MOTH SPECIES DIVERSITY 
USING A LIGHT TRAP IN A NORTHEASTERN SOFTWOOD 
FOREST A. W. Thomas and G. M. Thomas .................. 85 


A REVIEW OF THE PHLOXIPHAGA GROUP OF THE GENUS HELIO- 
THIS (NOCTUIDAE: HELIOTHENTINAE) WITH DESCRIPTION OF 
A NEW SPECIES David F: Hardwick... pa 106 


LIFE HISTORY OF PACHLIOPTA RHODIFER (PAPILIONIDAE: TROI- 
DINI) K. Veenakumari and Prashanth Mohanraj A EI 


RECENT INTRODUCTIONS OF RIPARIAN NOCTUID MOTHS FROM THE 
PALEARCTIC REGION TO NORTH AMERICA, WITH THE FIRST 
REPORT OF APAMEA UNANIMIS (HUBNER) (NOCTUIDAE: 
AMPHIPYRINAE) Kauri Mikkola and J. Donald Lafontaine 121 


SYSTEMATIC POSITION OF SIBATANIOZEPHYRUS AND DESCRIPTION 
OF A NEW SPECIES FROM TAIWAN (LYCAENIDAE: THECLINAE) 
Yu-Feng Hsu and Ming-Yao Lin ____ ss 


AVIAN PREDATION ON THE MONARCH BUTTERFLY, DANAUS 
PLEXIPPUS (NYMPHALIDAE: DANAINAE), AT A CALIFORNIA 
OVERWINTERING SITE Walter H. Sakai ... -) oa 148 


BEHAVIORAL RESPONSE OF MONARCH BUTTERFLIES (NYMPHALIDAE) 
TO DISTURBANCES IN THEIR HABITAT—A GROUP STARTLE 
RESPONSE? William H: Calvert 2 2 eee oe eer 


GENERAL NOTES 


The case of the missing H: Heliconius charithonia (L., 1767), not “ Heliconius 
charitonia (L., 1767). Andrew V. Z. Brower 2...) eee 166 


Native pierine butterfly (Pieridae) adapting to naturalized crucifer? A. V. 
Courant, A. E. Holbrook, E. D. Van der Reijden and F. S. Chew 168 


Implications of introduced garlic mustard (Alliaria petiolata) in the habitat 
of Pieris virginiensis (Pieridae) Aa POTte i o.eccccccnceeeecceeenceeeeeeeeeeeeeeee iyi! 


This paper meets the requirements of ANSI/NISO Z39.48-1992 (Permanence of Paper). 


— 


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544 Volume 48 1994 | Number 3 
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Cover illustration: The European tortoise-shell butterfly, Aglais urticae, flying over 
nettle, Urtica dioica. Original drawing by Hedvig Wael © @Ostern, Torsarudstubben 25, 
3430 Spikkestad, Norway. 


JouRNAL OF 


Tue LEPIDOPTERISTS’ SOCIETY 


Volume 48 1994 Number 3 


Journal of the Lepidopterists’ Society 
48(3), 1994, 173-179 


NECTAR SOURCE DIVERSITY AS AN INDICATOR OF 
HABITAT SUITABILITY FOR THE ENDANGERED 
UNCOMPAHGRE FRITILLARY, BOLORIA ACROCNEMA 
(NYMPHALIDAE) 


HuUGH B. BRITTEN AND LYNN RILEY 


Nevada Biodiversity Research Center, Department of Biology, 
University of Nevada, Reno, Nevada 89557, USA 


ABSTRACT. Pairwise comparisons between occupied and apparently suitable, yet 
unoccupied, Boloria acrocnema sites in the San Juan Mountains of southern Colorado 
indicated that occupied sites have a greater diversity of nectar source genera than un- 
occupied sites. This suggests a means of improving the efficiency of search efforts for 
additional populations of this endangered butterfly. 


Additional key words: habitat, Colorado, conservation, biological indicators. 


The Uncompahgre fritillary, Boloria acrocnema (Gall and Sperling) 
(Nymphalidae), was discovered on a high ridge near Uncompahgre 
Peak in the San Juan Mountains, Hinsdale County, Colorado in 1978 
(Gall & Sperling 1980). A second colony subsequently was found in 
1982 near Red Cloud Peak, approximately 16 km south of the type 
locality (Gall 1984a). Despite considerable effort throughout the 1980’s, 
U.S. Fish and Wildlife Service, Forest Service, and Bureau of Land 
Management personnel and others were unable to document the exis- 
tence of any additional colonies. Gall (1984b) estimated the total brood 
size of the Uncompahgre Peak colony at 650-750 individuals in 1980. 
The Red Cloud colony consisted of 1,000-1,500 individuals in 1982 
(Gall 1984b). Anecdotal accounts indicated that both known colonies 
were declining. 

Given the extreme endemicity of this butterfly and its apparent rarity, 
the species was considered for endangered status under the U.S. En- 
dangered Species Act in 1982 (Federal Register, Vol. 49, No. 100, P. 
2167). Work was begun in 1987 to characterize the habitat of the 
Uncompahgre fritillary, determine the status of the known populations, 


174 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


and discover any existing colonies near the known ones (Britten & 
Brussard 1992, Britten et al. 1994). Because the populations were de- 
clining, and because the odd-year brood of this biennial butterfly was 
apparently extirpated at its type locality in 1987 (Britten et al. 1994), 
the Uncompahgre fritillary was listed as endangered in 1991 (Federal 
Register, Vol. 56, No. 121, P. 28712). 

The 1993 draft recovery plan for the Uncompahgre fritillary stip- 
ulates that delisting of the butterfly will occur when stable colonies exist 
for 10 consecutive years at 10 or more locations. Because only small 
numbers (n<15) of butterflies were located at three additional sites 
(Brussard & Britten 1989), delisting will occur only with the discovery 
of a substantial number of new colonies. However, the search for col- 
onies is hampered by a number of difficulties. First, the known colonies 
exist at high elevations, approximately 4,000 m, in areas which generally 
are inaccessible by road. Thus, foot travel or high altitude helicopters 
are the only realistic means of access to potential habitat areas. Second, 
this butterfly’s flight season is short, lasting from early- to mid-July to- 
early August. In addition, weather conditions are unpredictable at high 
elevations. This diminishes further the duration of suitable flight con- 
ditions in which searches can be conducted. As poor flight conditions 
frequently occur during investigations, it is often impossible to elim- 
inate a site as potential habitat. The final difficulty in locating new 
colonies is that apparently suitable habitat is ubiquitous in the San Juan 
Mountains and nearby ranges. 

Larval B. acrocnema feed on snow willow, Salix nivalis (Booth) 
(Salicaceae) (Scott 1986), which occurs in fairly discrete patches from 
below tree-line to near the limit of vegetative growth in the San Juan 
Mountains. The two known colonies and other areas where the butterfly 
has been located, have approximately 15% snow willow cover, occur 
on northeast facing slopes with up to 45% grades, and contain snow 
fields and numerous flowers during the flight season (Brussard & Britten 
1989). These characteristics describe a very large portion of the alpine 
zone of the southern Rocky Mountains. All sites examined have similar 
habitat components, yet only a small fraction of them support B. ac- 
rocnema populations. Thus, an efficient method of habitat assessment 
is much needed. 

A number of studies have correlated adult butterfly resources, such 
as nectar supply, with population densities and individual movements 
(Gilbert & Singer 1975, Thomas & Singer 1987, Williams 1988). We 
examined data on nectar source inflorescences to test for a similar 
correlation between occupancy at potential B. acrocnema sites and 
nectar source abundance. Any such quantifiable differences between 


VOLUME 48, NUMBER 3 Ws 


occupied and unoccupied sites could then be used to establish search 
priorities among sites for this rare butterfly. 


MATERIALS AND METHODS 


Data were gathered during the 1988 flight season from the five 
occupied sites and from five unoccupied sites which were paired to the 
former on the basis of elevation, slope, aspect, geographical location, 
and percent cover of snow willow. Unoccupied sites were no more than 
200 m from the occupied sites with which they were paired. The 
unoccupied sites originally had been identified as good potential sites, 
due mostly to the presence of a patch of snow willow, and were surveyed 
for butterflies, along with a total of approximately 50 such sites, in the 
initial efforts to locate new colonies. The unoccupied sites were selected 
for comparison with the occupied sites because of their proximity to 
the latter. Nearness of paired sites is desirable for these comparisons, 
because it insures that unoccupied sites are as ecologically similar to 
ine occupied sites as possible. 

_Data on frequency of nectar source inflorescences by genus were 
determined using a line intercept method along habitat transects (Smith 
1980). This was accomplished by laying a 50 m tape measure across 
the potential habitat patch and tallying all flowering plants which came 
into contact with the tape measure (Britten 1991). Flowering plants are 
referred to as “nectar sources” if B. acrocnema was observed visiting 
the species during the flight season (Britten 1991). Transect length 
varied from 75 m to 150 m according to the length of the site, but the 
majority of sites were sampled with 100-m transects. Size of a site was 
not correlated with occupancy. Sites were characterized by frequency 
of inflorescences by genus (occurrence per meter) and the number of 
genera represented. 

The hypothesis that occupied sites are floristically more diverse 
than unoccupied sites was tested against a null hypothesis that unoc- 
cupied sites are equally or more floristically diverse than occupied sites. 
Thus, one-tailed statistical tests were appropriate for the rejection of 
the null hypothesis. 

The diversity of nectar source genera was compared among paired 
sites using N, the number of genera per site, and 1/D, the reciprocal 
of the Berger-Parker index of dominance (Magurran 1988). The Berger- 
Parker index indicates how dominant the most common taxon is within 
a site. Its reciprocal, therefore, gives an estimation of evenness. Taken 
together, taxon richness (N) and evenness (1/D) estimate site diversity. 
These data may not be normally distributed, and, thus, they require 
nonparametric analysis. The Wilcoxon signed ranks test was used to 


176 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


TABLE 1. Inflorescences present (+) or absent (—) on paired occupied (O) and un- 
occupied (U) Boloria acrocnema sites in the San Juan Mountains. 


Sites 


Uncompahgre Red Cloud A B C 


Genus O U O U O U 0) U O U 


Aster 

(Asteraceae) SF ze = + ar = + + + + 
Bistorta 

(Polygonaceae) — + = s Bb az at a au Bt 
Caltha 

(Ranunculaceae) — = = = es its x a a om 
Castilleja 

(Scrophulariaceae) + + + + + — + + + + 
Draba 

(Cruciferae) — = ae ee 2 os bs ny ia i) 
Dryas 

(Rosaceae) — = = = be = a as rH fe 
Hymenoxys 

(Asteraceae) + - _ == a pa hs es = ag 
Lomatium 

(Umbelliferae) — — a. = a! ot ae - ve Bt 
Mertensia 

(Boraginaceae) =F ~ - — - = zs as a of 
Penstemon 

(Scrophulariaceae) — _ — = = of =| an) » a 
Phlox 

(Polemoniaceae) + = - - - £ fe i = x 
Polygonum 

(Polygonaceae) + ~ + + = a a oak un = 
Potentilla 

(Rosaceae) + + =F + + + ar + + = 
Silene 

(Caryophyllaceae) = = + ae + + a ee = fh 
Trifolium 

(Fabaceae) + - + = = zal a es a Ps 
Zigadenus 

(Liliaceae) — = + = = - Lt B = oy 


compare N and 1/D between occupied and unoccupied sites. Paired 
sites were further compared using the frequencies of all flowers en- 
countered per meter of transect and the frequency of Potentilla (Ro- 
saceae) species encountered per meter of transect. Potentilla was chosen 
for this analysis because it was by far the most abundant flower on the 
study sites. These final between-site comparisons were made with a 
t-test, as nonparametric tests were not required (Sokal & Rohlf 1981). 


RESULTS 


The results of the analyses demonstrate that occupied B. acrocnema 
colony sites are more floristically diverse than nearby unoccupied sites. 
Flowers present at each site are indicated in Table 1. The number of 


VOLUME 48, NUMBER 3 ae 


TABLE 2. Comparison of nectar source diversity for paired occupied and unoccupied 
Boloria acrocnema colony sites. N is the number of nectar source genera represented on 
each site and 1/D is the reciprocal of the Berger-Parker index of dominance (Magurran 
1988). 


N 1/D 
Paired sites "Occupied —~Unoccupied Occupied Unoccupied 
Uncompahgre* 8 4 3.00 2.09 
Red Cloud 8 re) 1.70 Waa 
A 4 + 1.65 1.50 
B 8 4 3.10 1.20 
C ) 3 3.20 1.30 
P = 0.03 P = 0.04 
1-tailed Wilcoxon signed ranks tests 
* Type locality. 


genera in flower per site, N, was higher at occupied than unoccupied 
sites (P=0.03; Table 2). The reciprocal of the Berger-Parker index, 1/D, 
was higher at occupied sites (P=0.04; Table 2). The number of flowers 
encountered per meter was higher at occupied sites (P=0.04; Table 3). 
This may be, in part, because of the dominance of Potentilla species 
at most sites. The frequency of Potentilla species was higher at occupied 
sites (P=0.03; Table 3). Finally, Trifolium species (Fabaceae) occurred 
at four out of the five occupied sites, but at none of the unoccupied 
sites (Table 4). 


DISCUSSION 


The results of the present study are similar to those of Williams 
(1988) for the montane checkerspot Euphydryas gillettii (Barnes) 
(Nymphalidae) in the Rocky Mountains. Williams (1988) quantified 10 
habitat variables including topographic characteristics, larval host abun- 
dances, and nectar source abundances at 15 E. gillettii sites in Wyoming, 


TABLE 38. Comparison of flower density among paired occupied and unoccupied 
Boloria acrocnema sites. 


Flowers/meters Potentilla/meter 
Paired sites Occupied Unoccupied Occupied Unoccupied 
Uncompahgre* 0.16 0.15 0.05 0.07 
Red Cloud 0.57 0.19 0.33 0.10 
A 1.19 0.12 0.72 0.08 
B 1.67 0.20 0.46 0.16 
C 0.73 0.47 0.23 0.00 
P = 0.04 P = 0.03 


]-tailed t-tests 
* Type locality. 


IAs) JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


TABLE 4. Presence (+) or absence (—) of Trifolium species in flower at paired occupied 
and unoccupied Boloria acrocnema sites. 


Paired sites Occupied Unoccupied 


Uncompahgre 
Red Cloud 

A 

B 

Cc 


++ 144 


Montana, Idaho, and Alberta. Nectar source abundance was the only 
variable that significantly correlated with E. gillettii colony size (Wil- 
liams 1988). Likewise, the higher floristic diversity of occupied B. ac- 
rocnema sites relative to nearby unoccupied sites is probably the result 
of ecological correlations rather than direct causal mechanisms, such 
as relative nectar source availability. In other words, habitat charac- 
teristics which make a site more suitable to flowering plants also may 
do the same for B. acrocnema. Therefore, it is possible that some as 
yet unknown set of habitat variables determine habitat suitability for 
B. acrocnema and that the frequency of inflorescences is an ecological 
indicator of these favorable habitat sites. 

The search efforts of federal personnel up to this point have relied 
on three criteria for the identification of potential B. acrocnema colony 
sites: 1) the presence of snow willow, 2) northeastern aspects, and 8) 
approximate elevations greater than 3500 m. The results of the present 
study suggest that it would be prudent to include some assessment of 
flower frequency in the evaluation of potential habitat. This perhaps 
could be accomplished with a technique as simple as estimating the 
density of flowering Potentilla plants and noting the presence of Tri- 
folium on all investigated sites. 

High quality Boloria acrocnema habitat seems to be characterized 
by quantifiable parameters with respect to nectar source inflorescences. 
These habitat measures should not replace population studies, but they 
may be used to expand search efforts temporally. Thus, whereas current 
search efforts are possible only during the three week flight season, 
habitat assessment including nectar source evaluation could occur 
throughout the summer. Such inventories could be used to locate new 
colonies and prioritize sites for further monitoring during the flight 
season. 


ACKNOWLEDGMENTS 


We thank Peter Brussard and Richard Rust for reviewing early drafts of this work. 
Lawrence Gall and an anonymous reviewer offered many helpful criticisms of the manu- 
script. The Nevada Biodiversity Research Center provided partial support for this work. 


VOLUME 48, NUMBER 3 179 


LITERATURE CITED 


BRITTEN, H. B. 1991. The conservation biology of the Uncompahgre fritillary and the 
related northern dingy fritillary. Ph.D. Dissertation, Montana State University, Boze- 
man, Montana. 127 pp. 

BRITTEN, H. B. AND P. F. BRUSSARD. 1992. Genetic divergence and the Pleistocene 
history of the alpine butterflies Boloria improba (Nymphalidae) and the endangered 
Boloria acrocnema (Nymphalidae) in western North America. Canad. J. Zool. 70: 
539-548. 

BRITTEN, H. B., P. F. BRUSSARD, AND D. D. MurpHy. 1994. The pending extinction of 
the Uncompahgre fritillary butterfly. Conserv. Biol. 8:86—94. 

BRUSSARD, P. F. AND H. B. BRITTEN. 1989. The status of the Uncompahgre fritillary 
(Boloria acrocnema). Final report prepared for the U.S. Forest Service, Bureau of 
Land Management, and Fish and Wildlife Service. Montana State University, Boze- 
man, Montana. 47 pp. 

GALL, L. F. 1984a. Population structure and recommendations for conservation of the 
narrowly endemic alpine butterfly, Boloria acrocnema (Lepidoptera: Nymphalidae). 
Biol. Conserv. 28: 111-138. 

GALL, L. F. 1984b. The effects of capturing and marking on subsequent activity in 
Boloria acrocnema (Lepidoptera, Nymphalidae), with a comparison of different 
numerical models that estimate population size. Biol. Conserv. 28:189-154. 

GALL, L. F. AND F. A. H. SPERLING. 1980. A new high altitude species of Boloria from 
southwestern Colorado (Nymphalidae), with a discussion of phenetics and hierarchical 
decisions. J. Lepid. Soc. 43:230-252. 

GILBERT, L. A. AND M. C. SINGER. 1975. Butterfly ecology. Ann. Rev. Ecol. Syst. 6:365- 
397. 

MAGURRAN, A. 1988. Ecological diversity and its measurement. Princeton University 
Press. 179 pp. 

ScoTT, J. A. 1986. The butterflies of North America. Stanford University Press, Stanford. 
583 pp. 

SMITH, R. L. 1980. Ecology and field biology, third edition. Harper & Row, Publishers, 
New York. 835 pp. 

SOKAL, R. R. AND F. J. ROHLF. 1981. Biometry, second edition. W. H. Freeman & Co. 
859 pp. 

THOMAS, C. D. AND M. C. SINGER. 1987. Variation in host preference affects movement 
patterns within a butterfly population. Ecology 68:1262-1267. 

WILLIAMS, E. H. 1988. Habitat and range of Euphydryas gillettii (Nymphalidae). J. 
Lepid. Soc. 42:37—45. 


Received for publication 17 July 1998; revised and accepted 9 October 19938. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 180-189 


HIGH ANDEAN PRONOPHILINI FROM VENEZUELA: 
TWO NEW SPECIES OF DIAPHANOS 
(NYMPHALIDAE: SATYRINAE) 


ANGEL L. VILORIA P. 


Museo de Biologia, La Universidad del Zulia, Apartado 526, Maracaibo 4011, 
Zulia, Venezuela 


Abstract. Diaphanos fuscus, new species, and D. curvignathos, new species, are 
described and illustrated from the Cendé and Niquitao areas, respectively, in the States 
of Lara and Trujillo in the Andes of Venezuela. The two new species are compared with 
the type-species of the genus, D. huberi, which is known from the Cordillera de Mérida. 
A key for separating the three species is presented. Comments on the biology, habitat, 
and distribution of the new taxa are presented. 


Additional key words: Diaphanos fuscus, D. curvignathos, D. huberi, endemic spe- 
cies, paramo. 


The montane butterfly fauna of northern South America is excep- 
tionally diverse, especially the Satyrinae, which contains many endemic 
elements. Within the subfamily, the genera Paramo, Dangond, Re- 
donda, and Diaphanos are considered “‘relicts’’ (Adams & Bernard 1981, 
Adams 1985); they are restricted to high altitudes in the isolated paramos 
of the Sierra Nevada de Santa Marta (Colombia), Sierra de Perija (Co- 
lombia-Venezuela), and Cordillera de Mérida in Venezuela. 

Adams and Bernard (1981) described the genera Redonda and Dia- 
phanos from the highlands of the Mérida range, believing both to be 
monobasic. Diaphanos huberi Adams and Bernard is the most unusual 
satyrid of the tribe Pronophilini because of its small size, coloration 
(almost transparent), and other peculiar morphological features. 

During an inspection of the butterfly collection at the Museo del 
Instituto de Zoologia Agricola of the Universidad Central de Venezuela, 
I found a single worn male of a dark Diaphanos from Paramo del 
Jabon, Lara State, which was distinct from D. huberi. This led me to 
visit Fila de Los Nepes (near Paramo del Jabon) in August 1991, where 
I observed the butterfly in its natural habitat and collected additional 
specimens. Subsequently, J. Camacho of the Universidad del Zulia 
(MALUZ) showed me similar specimens from Paramo de Las Rosas, 
south of Los Nepes and nearer El Jabon, a place where this species is 
locally common. It became apparent that all these specimens repre- 
sented an undescribed taxon. I was interested in establishing the western 
distributional limit of this new Diaphanos, and during additional field 
work, I collected the genus in the Paramo de Ortiz, Trujillo State. The 
latter specimens are smaller in size and lighter in color. The genitalia 
are distinct, indicating that the specimens from Trujillo represent an 
additional undescribed species. In this paper I describe these two new 


VOLUME 48, NUMBER 3 181 


species and present life history observations. A key to the species based 
on wing coloration and distribution also is presented. 

Disposition of material. The holotypes and most of the paratypes 
are deposited in the Museo de Artropodos de la Facultad de Agronomia 
of La Universidad del Zulia (MALUZ) and in the Museo de Biologia 
de La Universidad del Zulia (MBLUZ), both in Maracaibo, Venezuela. 
Representative paratypes are deposited in the Museo del Instituto de 
Zoologia Agricola de la Universidad Central de Venezuela, Maracay 
(MIZA); The Natural History Museum, London, England (BMNH); and 
the private collection of Tomasz Pyrez, Warsaw, Poland (TPP). 


SYSTEMATICS 
Key to the Species of Diaphanos in the Venezuelan Andes 


1. Wings translucent, Mérida range Ui eens oceans UD). uber 
Wings not translucent; dorsally fuscous or dark brown wee ee 8 
2. Hindwing underside with one cream-white streak in cell A, (Cendé range) __. 
ee 2) | ee Siedler en ED SF USCHS, HS SP: 
Hindwing underside with two cream-white streaks in cell A, (Niquitao range) __ 
MOURN DD. Curvignathos, n- sp. 


Diaphanos fuscus Viloria, new species 


(Figs. 1, 2, 4, 6) 


Description. Eyes glabrous, dark brown, circled with reddish brown. Palpus twice as 
long as head, chestnut brown with dark fuscous hairs. Antenna to 0.40 length of costa; 
shaft orange-brown, sparsely scaled; club black, tapered (not as blunt as in D. huberi), 
2.5 to 3.0 times longer that broad, concave, comprising eleven segments. Wing venation 
as in D. huberi. Forewing tornus obtuse. Both wings rounded; dorsal surface covered with 
spindlelike scales that are dentate distally, broader distally (triangular shaped) in females. 
Marginal scales spinelike, each dentate at distal extreme; scales longer in female (Fig. 6). 

Male. Forewing length 17-21 mm (n = 28) (Fig. 1). Body dark fuscous, tip of abdomen 
lighter. Upperside ground color of wings dark fuscous, glossy, slightly lighter towards 
outer margins. Dorsal surface of wings hairy; hairs longer and more abundant in basal 
half, especially on hindwings. Underside ground color as on upperside, but lighter and 
more opaque. Hindwing underside with spindlelike cream-white streak parallel to veins, 
extending through discal cell and M, from base to near outer margin, interrupted by M,- 
M,; a series of narrow cream-white streaks parallel to the veins, from near discal cell to 
near outer margin, in cells Cu,, Cu,, and A,. Ventral surface of wing densely hairy. 
Genitalia as in Fig. 4. 

Female. Forewing length 16-19 mm (n = 8) (Fig. 2). Dorsal surface of body dark 
fuscous, ventral surface brown. Wings slightly narrower and forewing apex sharper than 
in male. Scales and hairs on both wing surfaces less dense than in male. Upperside ground 
color fuscous, darker at base, dusted with brown scales mainly in distal half of both wings. 
Underside ground color light brown, with golden sheen, dusted throughout with fuscous 
scales, more dense in basal one-third of both wings. Forewing underside with series of 
narrow and vestigial white streaks parallel to veins, from discal area to outer margin, in 
cells M, to Cu,; cell A, dusted with white scales near tornus. Hindwing underside with 
series of white streaks as in male, but less distinct. 

Variation. Wing maculation is variable in males. In a few specimens the forewing 
upperside is dusted with gray scales in the discal and postdiscal areas. The ground of 


182 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


VOLUME 48, NUMBER 3 183 


both the upper and underside may be olivaceous or brownish. In some individuals the 
distal one-fourth of the fore- and hindwing undersides have a chestnut tone. All streaks 
on the hindwing underside are highly variable. They may reach the outer margin or not; 
sometimes only vestigial streaks are present (except for the longer spindlelike one which 
is always well defined). The most strikingly marked individuals have an additional streak 
in cell M, and a speckling of cream-white scales near the streak in Ag. 

Variation among females is apparent on the underside of the wings where the ground 
color may be as light as cream-white. In this case it is difficult to distinguish the white 
streaks on both wings, except for the longer spindlelike streak of the hindwing. The most 
heavily marked individuals have additional narrow streaks in cell M, and Ag. 

Types. Holotype: male, Fila de Los Nepes, Paramo de Los Nepes, 2550 m, Lara State, 
Venezuela (9°45'N, 70°04'’W), 12 Aug. 1991, A. Viloria & J. Camacho (MALUZ). Paratypes: 
22 males, 8 females as follows: VENEZUELA: Lara State: 3 males, 4 females, same data 
as holotype (MALUZ); 1 male, Paramo del Jabon (9°30'N, 70°06’W), 3000 m, 20 Feb. 
1985, CEUM (MIZA); 18 males, 4 females, Paramo de Las Rosas, Municipio Moran (9°36'N, 
70°07'W), 3150 m, 20 Aug. 1991, J. Camacho (MIZA, BMNH, TPP, MALUZ). 

Etymology. The name fuscus refers to the dark coloration of this species. 


Comments on the Natural History. Diaphanos fuscus is locally com- 
mon in the paramos of Los Nepes, Las Rosas, and El Jabon, all of which 
belong to the massif of El Cendé (Fig. 7). El Cendé is considered one 
of the most isolated paramo units of the Venezuelan Andes (Simpson 
1971, Vuilleumier 1979). The species probably is restricted to this par- 
amo unit which includes the states of Lara and Trujillo. 

Northeastern slopes of the Cendé range (the Los Nepes area) are 
relatively arid; consequently, the paramo vegetation—with character- 
istic speletine composites—is developed from 2400 m, although there 
are some forest patches near La Fila (2500 m). Diaphanos fuscus 
flies only in paramo habitat and has been collected from 2550 to 3150 
m. : 

During a visit to Los Nepes in August 1991, the weather was mostly 
windy and foggy. No butterflies were seen until the clouds cleared 
bewteen 1140 and 1330 h. All individuals observed and collected were 
from a small open area surrounded by low shrubs (1.5 m tall) of Li- 
banothamnus (Asteraceae). The butterflies flew weakly, close to the 
grasses and over the shrubs, and never more than 2 m above the ground. 

A single female was observed resting and displaying on the flowering 
spike of a “‘palmiche” (Orthosanthus chimboracensis Bak., Iridaceae) 
in a supposed thermoregulatory posture exhibited by other paramo 
butterflies (Descimon 1986, Viloria unpublished). Two other females 


— 


Fics. 1-3. Upperside (left) and underside (right) of Diaphanos species. 1, D. fuscus, 
n. sp., female, same data as holotype; 2, D. fuscus, n. sp., holotype, male, Fila de Los 
Nepes, Paramo de Los Nepes, Lara State, Venezuela; 3, D. curvignathos, n. sp., holotype, 
male, Paramos de Ortiz, Trujillo State, Venezuela. 


184 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


4 -e) 


(CS Se eee 


It oot llooe / IM) 1009 h Siviv, 


AB COD Ay (Bo 3CeD A B 
D. curvignathos D. fuscus D. huberi 


Fics. 4-6. Male genitalia and wing scales of Diaphanos species. 4, Male genitalia of 
D. fuscus; 5, Male genitalia of D. curvignathos; 6A, Marginal scales in male; 6B, Dom- 
inant scales on wing surface of males; 6C, Marginal scales in females; 6D, Dominant 
scales on wing surface of females. 


flew from some palmiches when these plants were disturbed. Three 
females from Los Nepes remained alive for at least 30 hours following 
their capture. Each oviposited 4 to 15 eggs inside paper envelopes 
without attaching them to any surface. The most prolific female died 
still containing many eggs in its abdomen. 

The egg (Fig. 8) is spherical (1.0 mm in diameter), pale green, smooth, 
and free of cement or glue. Examples are deposited in MBLUZ. 

It is interesting that some females of Redonda from Paramo de Ortiz 
(Trujillo) had a similar but more violent oviposition behavior, releasing 
large numbers of eggs into paper envelopes immediately after their 
capture (Viloria personal observation). This may be an instinctive re- 
sponse to the stress of capture and handling. It also suggests that these 
butterflies may not place their eggs directly on the host, but may 


VOLUME 48, NUMBER 3 185 


broadcast them. Non-plant oviposition substrates have been reported 
previously for butterflies by Chew and Robbins (1984). In addition, 
DeVries (1987) reported that some Neotropical satyrid species expel 
their eggs while flying, so that eggs are dispersed over the plants or 
among the grasses on the ground, and this may be the case in D. fuscus. 
As for all Pronophilini, it is believed that a bambusoid grass is the 
host plant of Diaphanos fuscus. In my experience, bamboos are scarce 
in Los Nepes (but much more common in Las Rosas), and they grow 
mainly in the herbaceous strata under Libanothamnus shrubs. The only 
bamboo species recorded from Los Nepes is Rhipidocladum geminatum 
(McClure) (Clark & Londono 1991), but other species occur within the 
Cendé massif [e.g., Clarke (1989) reported Chusquea angustifolia (Sod- 
erstrom & C. Calderon) in Las Rosas]. Other dominant plant species in 
the highlands of Cendé are the grasses Agrostis and Calamagrostis 
(Poaceae) and the composites Espeletia, Espeletiopsis, Ruilopezia, and 
Hinterhubera (Asteraceae) (Vareschi 1970, Cuatrecasas 1979). The only 
pronophiline butterfly recorded in association with D. fuscus is an 
undescribed species of Redonda (Viloria & Pyrcz unpublished). 


Diaphanos curvignathos Viloria, new species 


(Figs. 3, 5, 6) 


Description. Eyes glabrous, dark brown. Palpus twice as long as head, light brown, 
with fuscous and chestnut hairs. Antenna to 0.4 length of costa; shaft orange-brown, 
almost without scales; club black, slightly more gradual than in D. fuscus, three times 
longer than broad, concave, comprising eleven segments. Venation similar to other species 
of Diaphanos. Forewing tornus obtuse; both wings rounded. Most of dorsal surface covered 
with suboval scales, longer on hindwing, broader and rounded in females. Some lighter, 
spindlelike scales, like those of D. fuscus male, dusted over entire wing surface. Marginal 
scales spinelike, dentate distally; longer in female (Fig. 6). 

Male. Forewing length 14-17 mm (n = 11) (Fig. 3). Body dark fuscous, slightly lighter 
on distal and ventral regions of abdomen. Upperside ground color of wings dark brown, 
glossy, lighter and reddish from distal region to outer margins. Dorsal surface of wings 
hairy; hairs longer and most abundant in basal one-half and anal region, especially long 
on hindwing. Basal one-half of forewing underside fuscous, becoming light brown in 
distal one-half. Hindwing underside light brown, becoming fuscous towards base and 
center of wing. Discal cell almost entirely cream-white; a series of narrow cream-white 
steaks parallel to veins, from discal cell to outer margin, in cells M, to A, (two streaks in 
latter). Ventral surface of wings densely hairy. Genitalia as in Fig. 5. 

Female. Forewing length 16 mm (n = 1). Essentially as described for male. 

Variation. Variation among the specimens studied is slight. In some males the upperside 
ground color of the distal two-thirds of the wings is ocherous rather than reddish, and 
the cream-white scales in the discal area and streaks on the hindwing underside are pale 
yellow. In darker individuals, the streaks are reduced but still distinct. In lighter indi- 
viduals, there are cream-white scales in the costal region of the hindwing underside. 

Types. Holotype: male, Paramo de Ortiz, 2850-3100 m, Trujillo State, Venezuela 
(9°13'N, 70°24'W), 12 Sept. 1991, A. Viloria & E. Moscé (MBLUZ). Paratypes: 10 males, 
1 female as follows: VENEZUELA: Trujillo State: 5 males, same data as holotype (BMNH, 
MBLUZ, TPP); 5 males, 1 female, Paramo de Ortiz, 2900-3000 m, 7 Jan. 1992, A. Viloria 
& J. Camacho (MALUZ). 

Etymology. The specific epithet curvignathos is a composite of the Latin curvus (= 


186 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


14 LAKE MARACAIBO 
& 
4 Zulia State 


Tachira “State. 


j ™~ Zulia State 


Merida State ———~. 


awis auf 


C 
—_gawis 007 


TRUJILLO 
e 


\ 
\ 


1 CENDE RANGE 
Land above 3000m 2 NIQUITAO RANGE 
@ D. curvignathos 3 SANTO DOMINGO RANGE 
@ D. fuscus 4 LA CULATA RANGE 
© D. huberi 5 MERIDA RANGE Q_ 19 20 30 40 50 7 


KM 


Fic. 7. Geographical distribution of the Venezuelan highlands and distribution of the 
three species of Diaphanos. 


curved) and the Greek gnathos (= mandible, also the name of the subuncal structure in 
the male genitalia), alluding to the hooked subunci of this species. 


Comments on distribution and biology. Diaphanos curvignathos 
occurs locally in Paramo de Ortiz, but is rare. It is possible that its 
distribution includes all of the paramos connected to the Niquitao 
massif, one of which is Ortiz. The latter range contains the highest 
mountains of Trujillo State, reaching 4006 m at Cerro La Teta. 

Because of the orographic connection between the Merida and Ni- 
quitao mountains and the fact that D. huberi ranges between 3400 and 
4000 m in Merida, it is possible that D. curvignathos and D. huberi 
are parapatric in the Niquitao range. However, D. huberi has been 
recorded only in the Cordillera de Mérida, and D. curvignathos is 
unknown from that range. 

Owing to the reduced area of paramo above 3400 m in Niquitao, it 
is unlikely that any other species ef Diaphanos occupies the uppermost 
level of these mountains, but further exploration is required. 

All specimens but one, which was collected on my first visit to Paramo 
de Ortiz, were collected within a few minutes at midday, flying over 


VOLUME 48, NUMBER 3 : 187 


1mm 


Fic. 8. Eggs of Diaphanos species. A, D. fuscus, B, D. curvignathos. 


a small area of grasses in a bend in the road from Trujillo to Bocono. 
This visit was during the wet season and many plants, especially com- 
posites, were in flower. During a second visit in January (dry season), 
it was sunny but adult butterflies were rare. The butterflies were flying 
over the grasses and resting on the ground. The only female was taken 
resting on the dirt road at 1630 h, and it expelled one egg immediately 
upon capture. The egg (Fig. 8) was spherical (0.8 mm in diameter), 
pale green, vertically ribbed, and free of cement or glue. It is preserved 
and deposited in MBLUZ. 

Diaphanos curvignathos has a relatively slow flight. It occurs only 
in paramo habitat between 2850 and 3100 m. The potential foodplant 
is an undetermined bamboo species (probably Chusquea) which is lo- 
cally abundant in Paramo de Ortiz, especially near streams. 

Other satyrids associated with D. curvignathos in Ortiz are Alto- 
pedaliodes albonotata (Godman) and an undescribed form of Redonda 
empetrus, all of which are sympatric. 


DISCUSSION 


There is sufficient evidence to conclude that Diaphanos huberi, D. 
fuscus, and D. curvignathos are three distinct species rather than sub- 
species. They differ in size, coloration, and genital structure. In general, 
D. fuscus is the largest and D. huberi the smallest. Differences in wing 
color patterns allow separation of the species following the key. 

Diaphanos fuscus and D. curvignathos are most similar in external 
appearance, but genitalia suggest a closer relationship between D. cur- 
vignathos and D. huberi, both of which have well developed subunci. 
In contrast, the subuncus of D. fuscus is atrophied. Diaphanos huberi 
has a shorter saccus than the other two species. The valvae and aedeagus 


188 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


are distinct in all three taxa, as are the shape of the wing scales (Fig. 
6). 

Apparently, each species is restricted to a small area of paramo, which 
may be the result of orographic and geographic isolation (Fig. 7). 
Diaphanos fuscus lives in the Cendé region whose paramos are quite 
isolated from the others in Venezuela. Diaphanos curvignathos lives in 
the highlands of Niquitao which are connected to those of the Cordillera 
de Mérida where D. huberi lives. 

Diaphanos huberi has not been found in Niquitao, nor D. curvig- 
nathos in Mérida. Therefore, it appears that there are other ecological 
factors limiting their distributions. Although these two species inhabit 
only open lands of the paramos, D. huberi is restricted to the uppermost 
level between 3600 and 4000 m. 


ACKNOWLEDGMENTS 


I am indebted to Jess Camacho, Rosanna Calchi (Universidad del Zulia), Edwin Mosco 
(Maracaibo), and Pedro Mora, Jr. (Trujillo) for their valuable assistance with fieldwork. 
I thank J. DeMarmels and L. D. Otero of Maracay for allowing me access to the butterfly 
collections of MIZA. DeMarmels also offered hospitality and help in Maracay. Pedro 
Mora, Sr., and his family in Las Adjuntas (Trujillo), and Juan Suarez and his family in 
La Aguada de Arenales (Lara) are unforgettable frineds whose warm hospitality I ac- 
knowledge. Many thanks to John Moody (Maracaibo), Michael Adams (Blandford, En- 
gland), Gerardo Lamas (Lima, Pert), and an anonymous referee for reviewing the original 
manuscript. I also thank José Moscé for encouraging me to continue working on the 
Venzuelan fauna, and for his constant help. Financial support for the fieldwork was 
provided by a grant from the Consejo de Desarrollo Cientifico y Humanistico to Magally 
Quirés, Director of MALUZ, and by MBLUZ. 


LITERATURE CITED 


ADAMS, M. J. 1985. Speciation in the pronophiline butterflies (Satyridae) of the northern 
Andes. J. Res. Lepid. Suppl. 1:33—49. 

ADAMS, M. J. & G.I. BERNARD. 1981. Pronophiline butterflies (Satyride) of the Cordillera 
de Mérida, Venezuela. Zool. J. Linn. Soc. 71:3438-372. 

CHEW, F. S. & R. K. Rospins. 1984. Egg-laying in butterflies. Symp. Roy. Entomol. 
Soc. London 11:65-79. 

CLARK, L. G. 1989. Systematics of Chusquea Section Swallenochloa, Section Verticil- 
latae, Section Serpentes, and Section Longifoliae (Poaceae: Bambusoideae). Syst. Bot. 
Monogr. 27:1-127. 

CuLaRK, L. G. & X. LONDONO. 1991. A new species and new section of Rhipidocladum 
(Poaceae: Bambusoideae). Am. J. Bot. 78:1260-1279. 

CUATRECASAS, J. 1979. Comparacion fitogeografica de paramo entre varias cordilleras, 
pp. 89-99. In Salgado-Labouriau, M.L. (ed.), El medio ambiente paramo. Centro de 
Estudios Avanzados del IVIC, Caracas. 237 pp. 

DESCIMON, H. 1986. Origins of lepidopteran faunas in the high tropical Andes, pp. 
500-532. In Montasterio, M. & F. Vuilleumier (eds.), High altitude tropical bioge- 
ography. Oxford Univ. Press, New York. x + 640 pp. 

DEVRIES, P. J. 1987. The butterflies of Costa Rica and their natural history. Papilionidae, 
Pieridae, Nymphalidae. Princeton Univ. Press, New Jersey. xxii + 326 pp. 

SIMPSON, B. B. 1971. Pleistocene changes in the fauna and flora of South America. 
Science 173:771-780. 


VOLUME 48, NUMBER 3 189 


VARESCHI, V. 1970. Flora de los paramos. Talleres Graficos Universitarios, Mérida. 429 


Pp. 
VUILLEUMIER, F. 1979. Comparacion y evolucion de las comunidades de aves de paramo 
y puna, pp. 181-205. In Salgado-Labouriau, M. L. (ed.), E] medio ambiente paramo. 


Centro de Estudios Avanzados del IVIC, Caracas. 237 pp. 


Received for publication 14 November 1992; revised and accepted 20 November 19938. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 190-198 


CONTINUED INTERSPECIFIC HYBRIDIZATION BETWEEN 
LIMENITIS (BASILARCHIA) ARTHEMIS ASTYANAX 
AND L. (B.) ARCHIPPUS IN THE 
SOUTHEASTERN U.S. (NYMPHALIDAE) 


AUSTIN P. PLATT 


Department of Biological Sciences, University of Maryland Baltimore County, 
Catonsville, Maryland 21228, USA 


AND 


JAMES R. MAUDSLEY 
400 University Circle, Athens, Georgia 30605, USA 


ABSTRACT. Interspecific hybridization between North American admiral butterflies 
is briefly reviewed and updated. Records of 77 wild male F, hybrids between the viceroy, 
Limenitis archippus subspecies, and other nearctic Limenitis are noted. Among these 
are six new records of the hybrid form “rubidus” Strecker (L. archippus x L. arthemis 
astyanax), including three each from two localities, one in Athens, Georgia and the other 
in northern Florida. Four of these new hybrids were reared from eggs and larvae wild- 
collected from willows (Salix spp.). Topics reviewed include 1) the present status of viceroy 
mimicry, and 2) the intense hybridization between phenotypically variable populations 
of L. archippus and L. arthemis astyanax observed recently in the Georgia/Florida region. 
Ecological, phenotypic, and behavioral reasons underlying this phenomenon are consid- 
ered. 


Additional key words: admirals, L. archippus floridensis, hybrid “rubidus,” mimicry, 
selection. 


Interspecific hybridization among North American Limenitis is well- 
known and has received much attention in the literature (see reviews 
by Platt et al. 1978, Platt 1983, Ritland 1990). Of particular interest to 
both lepidopterists and biologists are those crosses involving the viceroy, 
L. archippus (Cramer), a known mimic of various danaine models, and 
the other congeneric admirals with which L. archippus is broadly sym- 
patric. These include butterflies of the eastern L. arthemis-astyanax 
complex (Platt & Brower 1968, Platt 1975, 1987a, 1994) and species of 
the two western complexes, L. lorquini Boisduval (Perkins & Perkins 
1966, Gage 1970, Perkins & Gage 1970) and L. weidemeyerii Edwards 
(Cross 1986, 1937, Perkins & Perkins 1967, Simpson & Pettus 1976). 
Such crosses involving L. archippus and its congeners yield phenotyp- 
ically intermediate F, males, which are somewhat variable in terms of 
their relative light (orange) or dark (brownish-black) ground coloration, 
and also their partial postmedial white banding. 

These naturally occurring hybrid forms have been variously named 
(see below), depending upon the species (or subspecies) involved. These 
interspecific hybrids are quite rare, usually occurring as from one to 
several specimens in widely distributed, but geographically isolated, 


VOLUME 48, NUMBER 3 191 


localities. Some of these locales are referred to by collectors as hybrid 
“hot spots.” In such areas, low levels of interspecific cross-breeding take 
place over a number of years (and insect generations) in the same 
locality. Such wild hybrids presumably result from “stray” matings 
between individuals belonging to the two different taxa involved [e.g., 
1) L. arthemis arthemis x L. archippus, 2) L. arthemis astyanax x 
L. archippus, 3) L. weidemeyerii x L. archippus, and 4) L. lorquini 
x L. archippus]. Including the specimens to be reported in this paper, 
a total of 77 published and unpublished records of such interspecific 
hybrids presently are known to the senior author. The most recent of 
these records are from south-central Louisiana in September 1991 (Kemp 
1991) and southwestern Kentucky in September 1993 (Covell 1994, 
pers. comm.). These breakdown as follows: 


1) hybr. “arthechippus” Scudder = 11 (including the type specimen 
of hybr. “rubrofasechippus’’ Gunder) 

2) hybr. “‘rubidus’’ Strecker = 45 

3) hybr. “weidechippus” Cross = 12 

4) hybrid unnamed (L. lorquini x L. archippus) = 9 


All of these wild-caught specimens are males. However, a single het- 
erotic female specimen of hybr. “rubidus’’ has been lab-reared from 
Maryland strains (Platt & Harrison 1994). Similar female morphs occur 
in laboratory backcrosses involving the F, hybrid males as well (Platt 
1975, Platt et al. 1978). 


Hybrid “rubidus’’ Strecker 


Particularly well known are the F, hybrids between the two mimetic 
butterflies, L. arthemis astyanax and L. archippus subspecies. Such 
hybrids are broadly distributed from Arizona and New Mexico across 
the mid-western U. S. into New England and south to Florida. The two 
parent insects belong to two separate well-known mimicry complexes, 
and themselves are very different in appearance (Figs. 1 & 4). Thus, 
the morphologically intermediate F, hybrids represent a complete 
breakdown of both mimetic patterns (Fig. 3), and they quite likely are 
at a selective disadvantage from predation (by birds), when compared 
to either of the parental types. Because the known wild F, hybrids all 
are males, they most certainly are at a mating disadvantage as well. 
The hybrid morphs are not known to persist in nature, and backcross 
specimens at present are known only from laboratory crosses (Platt 
1975, 1988, Platt et al. 1978). 

Presently, little is known regarding courtship behavior in nearctic 
admirals. Possibly visual, tactile, and pheromonal cues are important 
to the insects, as has been shown for closely related palearctic species 


OZ JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


(Lederer 1960). Both parental species possess distinctive apical mor- 
phology of the male valvae (Platt et al. 1970), but apparently this pre- 
copulatory reproductive isolating mechanism is not perfect. The hybrids 
possess intermediate genitalic morphology. 

Results of laboratory hybridization studies (Platt 1975, 1983) reveal 
that some degree of inter-fertility is possible in reciprocal crosses, al- 
though a greater number of the few interspecific pairings encountered 
in nature involve female L. arthemis astyanax and male L. archippus 
archippus (Klots 1959, Ritland 1990). Apparently, the specialized elon- 
gate, sickle-shaped valval tips of L. archippus subspecies are efficient 
mating structures which may play an important functional role in 
interspecific hybridization among the nearctic Limenitis. However, 
recent observations by Covell (1994) in southwestern Kentucky and by 
S. Mertens (pers. comm.) in north central Wisconsin, demonstrate that 
the reciprocal crosses involving female L. archippus archippus X male 
L. arthemis astyanax and L. arthemis arthemis sometimes do take 
place in nature in the following article (Covell 1994:199). 


Present Status of Viceroy Mimicry 


Recently, the queen/Florida viceroy relationship (and hence, by in- 
ference, the monarch/viceroy relationship as well) has been shown to 
be a Miillerian one, rather than a Batesian one, as classically believed 
(Brower 1992, Ritland 1991, Ritland & Brower 199la, 1991b, Vane- 
Wright 1991, Walker 1991). These findings confirm those of J. V. Z. 
Brower (1958a, 1958b) and Platt et al. (1971) that certain avian pred- 
ators (scrub jays, Aphelocoma coerulescens coerulescens Bosc. and blue 
jays, Cyanocitta cristata bromia Oberholser; Corvidae) find viceroy 
butterflies unpalatable in caged experimental situations. 


RESULTS 


This paper reports the collecting of six additional wild “rubidus” 
hybrids, three each from Clarke Co., Georgia and Columbia Co., Flor- 
ida. All were taken between 1973 and 1986 by J. R. M. The Athens 
records probably involve the nominate subspecies, L. archippus ar- 
chippus, whereas those from Florida most likely involve the southern 
chocolate-brown subspecies, L. archippus floridensis Strecker, or L. 
archippus/floridensis intergrade forms, which are prevalent in north- 
ern Florida and southeastern Georgia. Four of these records represent 
specimens reared from wild-collected eggs and larvae found on willows 
(Salix spp.; Salicaceae). These records (presented chronologically) are 
as follows: | 

1) A worn adult male was collected at Belmont Road and Shoal 


VOLUME 48, NUMBER 3 193 


Creek in Athens (Clarke Co.), Georgia on 19 September 1973. The 
willow thicket was part of an ecotone habitat between the wooded 
bottomlands along Shoal Creek and an open cow pasture. 

2) A fresh adult male was captured at the ‘““Beaver Pond” site in 
Athens, on 17 May 1984. This area consists of a two- to three-acre beaver 
pond surrounded on three sides by open “old field” habitat, and by a 
mature pine plantation. The specimen was taken in a willow thicket 
between the beaver pond and the pine plantation. 

3) A third hybrid male eclosed on 1 August 1984 from a single larva 
collected several weeks earlier on Salix nigra Marsh at the Athens 
“Beaver Pond”’ site. 

4) Finally, three other male hybrids, all dark morphs of “rubidus,”’ 
were among 12 admirals reared from a group of 11 young larvae and 
two eggs collected along a 30 m stand of willow shrubs (believed to be 
Salix caroliniana Michx.) between a service road and a cypress swamp, 
located just north of the entrance to O’leno State Park (Columbia Co.), 
Florida on 11 July 1986. At the time, the willow leaves showed few 
signs of larval feeding, and only one freshly eclosed light orange female 
L. archippus archippus was seen flying in the area. 

_ This collection of eggs and larvae was made between summer broods 
of the adult admirals. Only 12 butterflies were reared, since one of the 
eggs failed to hatch. These insects were lab-reared at room temperature 
on Salix nigra, and eclosed between 19 July and 5 August 1986. Eclosion 
dates for the three hybrids were 19, 28, & 29 July. Also, reared in this 
sample were seven L. arthemis astyanax (4 males & 3 females) and 
two male L. archippus floridensis. All except one male L. arthemis 
astyanax represent the iridescent greenish form “viridis” Strecker. 


Intense Hybridization in the Florida/Georgia 
Subspecific ““Suture-zone”’ 


Assuming that all like butterflies were siblings, the 12 individuals 
must represent eggs laid by at least three different wild females. The 
last three hybrids well may involve the dark brown Florida subspecies 
of L. archippus (Fig. 2), although specimens seen and taken at this 
locality represent intraspecific intergrades, and vary greatly in ground 
color (ranging from bright orange to orange-brown to brown). This 
region, of course, represents an area of intergradation between the 
typical orange colored eastern viceroy and the darker peninsular sub- 
species L. archippus floridensis (Remington 1958, 1968). 

Ritland (1990) reports seven additional hybr. “rubidus’” records from 
Georgia and northern Florida obtained in 1986 and 1987. Five of his 
records, likewise, were obtained from wild-collected larvae taken on 
Salix caroliniana. There are, in addition, two earlier records of hybr. 


194 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


“rubidus’’ from this region (Platt et al. 1978). All 13 of the wild “rub- 
idus’ hybrids recently collected by J. R. M. & D. B. R. from this area 
represent progeny of either spring or early summer interspecific matings 
of the parental butterflies. Further north, and in western regions of the 
country, the majority of such crosses seem to occur in the late summer 
or fall (Platt 1987b). According to Ritland (1990), L. arthemis astyanax 
is absent from local habitats where L. archippus archippus and L. 
archippus floridensis fly during the late summer and fall seasons. 

Ritland (1990) discussed possible reasons for this cross-breeding. They 
include the following: 

1) The existence of the subspecific intergrade zone between L. ar- 
chippus archippus and L. archippus floridensis in southern Georgia 
and northern Florida, which creates greater genetic and phenotypic 
variability within populations of this often phenotypically uniform spe- 
cies. This greater genetic diversity within these viceroy populations 
may contribute to more labile female mate-choice. 

2) The general scarcity of L. arthemis astyanax in many regions of 
Florida, at least relative to L. archippus archippus, and the utilization 
of similar microhabitats by both species in the southeastern U.S., es- 
pecially during the early part of the year (April-June). [However, in 
northern Florida, L. arthemis astyanax populations can be reasonably 
common, locally.] 

3) In this area L. arthemis astyanax switches to greater dependence 
on salicaceous foodplants, since its main rosaceous foodplants, such as 
Prunus serotina Ehrh., become restricted to dry woodland habitats, 
and generally do not occur in the moist open meadow habitats, usually 
frequented only by viceroys. 


DISCUSSION 


Our present studies reveal the existence of phenotypic intergradation 
between L. archippus archippus and L. archippus floridensis in Georgia 
and northern Florida, as reported previously by Remington (1958, 1968). 
In fact, viceroy populations as far north as Athens, Georgia show ten- 
dencies toward this phenotypic blending with the darker southern sub- 
species. Such individuals usually are a darker orange-brown color than 
that of nominate L. archippus. Many of these specimens have forewings 
that are a shade darker than their hindwings, and some even possess 
ventral hindwing basal orange spots, like those characteristic of L. 
arthemis astyanax (Fig. 1). This last trait appears in L. archippus 
archippus because of differences in hues of the ventral hindwing ground 
color and of the basal spots themselves. Laboratory crosses between the 
two viceroy subspecies clearly show that the appearance of these ventral 


VOLUME 48, NUMBER 3 ; 195 


DORSAL 


VENTRAL orn 


Fics. 1-4. Specimens of the different forms of Limenitis from the southeastern U. S. 
1) L. archippus archippus, no. 84-1, Athens (Clarke Co.) Georgia, Aug. 1984, J. R. M.; 
2) L. archippus floridensis, no. 86-7, eclosed 3 Aug. 1986; 3) hybr. “rubidus’’, no. 86- 
11, eclosed 23 July 1986; & 4) L. arthemis astyanax, no. 86-3, eclosed 27 July 1986. All 
four specimens are males, reared on Salix sp. Specimens 2—4 reared from eggs and larvae 
collected on Salix sp. near O’leno State Park (Columbia Co.), Florida, 11 July 1986, by 
J. R. M. All specimens deposited in the insect collection at U.M.B.C. 


basal spots can occur without any cross-breeding involving viceroys and 
red-spotted purples. Thus, in itself, this trait among viceroys is not an 
indicator of such interspecific genetic introgression. 

Although throughout much of Florida L. arthemis astyanax seems 
to be relatively scarce compared to L. archippus, this does not appear 
to be the case in the O’leno State Park region of northern Florida. This 
area includes mesic habitat laced with ribbons of wet bottomlands and 
sink holes. These wetter low areas usually are wooded. The higher, 
more dry uplands form a patchwork of open fields, farmlands, and 
islands of either pines (Pinus spp.; Pinaceae) or oaks (Quercus spp.; 
Fagaceae). Within the more open pine areas and throughout the old 
fields, wild cherries and wild plums (Prunus spp.; Rosaceae) are among 


196 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


the most common successional tree species. Willows (Salix spp.) are 
much less common and are restricted to the wettest locales. Thus, L. 
arthemis astyanax usually is common and widespread around O’leno 
State Park, whereas L. archippus (both subspecies and their intergrades) 
is only locally abundant. Consequently, the interweaving of the two 
habitats quite likely plays the most important role in bringing the 
two species together in this area. The greater utilization of salicaceous 
foodplants by L. arthemis astyanax larvae has not been demonstrated, 
and, in fact, seems less likely to us as an important mechanism for 
bringing the two species together. It is true, however, that virtually all 
hybr. “rubidus” have been collected in rather open moist field or eco- 
tonal areas, having willows and water nearby. Such localities generally 
are considered to be viceroy habitats. 

An additional contributing factor to finding so many hybrids in such 
a short time period may be the increased intensity of collecting wild 
admiral larvae. This by itself has yielded nine (69%) of the 13 hybrids 
recently taken. Nearly all previous specimens were collected with hand- 
nets, following “random” encounters with collectors. At the same time, 
such rare insects are eagerly sought, and are collected whenever possible. 
In other words, the frequency of these cross-matings may not be just 
a recent phenomenon, but may have been occurring all along over a 
prolonged period of time. Such crosses simply may be an indication of 
the extremely close affinities between L. arthemis astyanax and L. 
archippus subspecies, as belied by their very similar developmental 
stages. It is likely that the red-spotted purple (L. arthemis astyanax) 
has evolved as a mimetic form from L. arthemis arthemis in compar- 
atively recent times. Thus, it may have spread southward into the 
vicinity of northern Florida very recently (geologically speaking) as 
well. 

Four of the hybr. “‘rubidus” specimens collected by J. R. M. are in 
his personal collection. The other two hybr. “rubidus’’ and the re- 
maining Florida specimens are in the U.M.B.C. insect collection. 


CONCLUSIONS 


Evidently, both the Beaver Pond site in Athens, Georgia, and the 
O’leno State Park locality in northern Florida represent the interspecific 
“hot spot” locales referred to earlier in this paper. In such habitats, 
ecological conditions including temperature, light, moisture, and food 
plant types and distributions, together with the relative abundances of 
the two parental species (with one being prevalent, but the other much 
scarcer) produce conditions leading to “microsympatry” at certain times 
of the year (Ritland pers. comm.). As long as such conditions persist, 


VOLUME 48, NUMBER 3 197 


elevated levels of hybridization can continue to occur at these specific 
locales. 


ACKNOWLEDGMEMTS 


The senior author thanks Connie Janni of Cleveland, Ohio, for initially bringing the 
Florida hybrids reared by J. R. M. to his attention. Two reviewers, R. C. Lederhouse and 
D. B. Ritland, provided valuable comments which greatly improved the manuscript. We 
also thank D. Flaim of UMBC for his worthwhile suggestions and for assistance with the 
manuscript preparation. G. C. Ford, Graphics Illustrator, U.M.B.C., prepared the plate 
(Figs. 1-4), 


LITERATURE CITED 


BROWER, J. V. Z. 1958a. Experimental studies of mimicry in some North American 
butterflies. I. The monarch, Danaus plexippus and viceroy, Limenitis archippus. 
Evolution 12:32—47. 

1958b. Experimental studies of mimicry in some North American butterflies. 
Ill. Danaus gilippus berenice and Limenitis archippus floridensis. Evolution 12: 
273-285. 

BROWER, L. P. 1992. The current status of butterfly royalty: monarchs and viceroys 
revisited. Terra 30(4):4—-15. 

COVELL, C. V., JR. 1994. Field observations of matings between female Limenitis 
archippus and male L. arthemis subspecies (Nymphalidae). J. Lepid. Soc. 48:199- 
204. 

Cross, F. C. 1936. (No title). Hobbies 41:112. 

. 1937. Butterflies of Colorado. Proc. Colo. Mus. Nat. Hist. 16:3-28. 

GaGE, E. V. 1970. A record of a naturally occurring Limenitis hybrid (Nymphalidae) 
J. Lepid. Soc. 24:270. 

Kemp, J. M. 1991. My “rubidus.” News Lepid. Soc. 6:82. 

Kiots, A. B. 1959. A mixed mating of two species of Limenitis Fabricius (Lepidoptera, 
Nymphalidae). J. New York Entomol. Soc. 67:20. 

LEDERER, V. G. 1960. Verhattensweissen der imagines und der entwicklungsstadien 
von Limenitis camilla camilla L. (Lep., Nymphalidae). A. Tierpsychol. 17:521-546. 

PERKINS, E. M. & E. V. GAGE. 1970. On the occurrence of Limenitis archippus x L. 
lorquini hybrids (Nymphalidae). J. Res. Lepid. 9:223-226. 

PERKINS, E. M., JR. & S. F. PERKINS. 1966. A review of the Limenitis lorquini complex. 
J. Lepid. Soc. 20:172-176. 

PERKINS, S. F. & E. M. PERKINS, JR. 1967. Revision of the Limenitis weidemeyerii 
complex, with description of new subspecies (Nymphalidae). J. Lepid. Soc. 21:213- 
234. 

PLATT, A. P. 1975. Monomorphic mimicry in nearctic Limenitis butterflies: experi- 
mental hybridization of the L. arthemis-astyanax complex with L. archippus. Evo- 
lution 29:120-141. 

1983. Evolution of North American admiral butterflies (Limenitis: Nymphal- 

idae). Bull. Entomol. Soc. Am. 29:10-22. 

1987a. Banded admirals from western Maryland; Analysis of the Limenitis 

(Basilarchia) arthemis-astyanax complex (Lepidoptera: Nymphalidae) at Green Ridge 

State Forest. Proc. Entomol. Soc. Wash. 89:633-645. 

1987b. Recent observations on North American admirals. Maryland Entomol. 

3:18-20. 

1994. Mimicry among butterflies: The subtleties of selection. Audubon Nat. 
News 20:8-10. 

PLATT, A. P. & L. P. BROWER. 1968. Mimetic versus descriptive coloration in inter- 
grading populations of Limenitis arthemis and astyanax butterflies. Evolution 22: 
699-718. 


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PLATT, A. P. & S. J. HARRISON. 1994. First record of an heterotic, adult female hybrid 
Limenitis (Basilarchia) “rubidus’’ (Strecker) (Lepidoptera: Nymphalidae). Entomol. 
News 105:33-38. 

PLATT, A. P., R. P. COPPINGER, & L. P. BROWER. 1971. Demonstration of the selective 
advantage of mimetic Limenitis butterflies presented to caged avian predators. Evo- 
lution 25:692-701. 

PLATT, A. P., S. D. FREARSON, & P. N. GRAVES. 1970. Statistical comparison of valval 
structure within and between populations of North American Limenitis (Nymphal- 
idae). Canad. Entomol. 102:513-5388. 

PLATT, A. P., G. W. RAWSON, & G. BALOGH. 1978. Interspecific hybridization involving 
Limenitis archippus and its congeneric species (Nymphalidae). J. Lepid. Soc. 32: 
289-303. 

REMINGTON, C. L. 1958. Genetics of populations of Lepidoptera. Proc. Tenth Int. Congr. 
Entomol. 2:787-805. 

. 1968. Suture-zones of hybrid interaction between recently joined biotas. Evol. 
Biol. 2:321—-428. 

RITLAND, D. B. 1990. Localized interspecific hybridization between mimetic Limenitis 
butterflies (Nymphalidae) in Florida. J. Lepid. Soc. 44:163-178. 

1991. Revising a classic butterfly mimicry scenario: Demonstration of Miullerian 
mimicry between Florida viceroys (Limenitis archippus floridensis) and queens 
(Danaus gilippus berenice). Evolution 45:918-934. 

RITLAND, D. B. & L. P. BROWER. 1991a. The viceroy is not a Batesian mimic. Nature 
350:497-498. 

1991b. A reassessment of the mimicry relationship among viceroys, queens, 
and monarchs in Florida, pp. 129-139. In Malcomb, S. B. & M. P. Zalulli (eds.), 
Biology and conservation of the monarch butterfly. Sci. Sers. 38, Los Angeles Co. 
Nat. Hist. Mus., Los Angeles, CA. 

Simpson, R. G. & D. PeTTus. 1976. Records of Limenitis hybrids in Colorado. J. Res. 
Lepid. 15:163-168. 

VANE-WRIGHT, R.I. 1991. A case of self-deception. Nature 350:460-461. 

WALKER, T. 1991. Butterflies and bad taste—rethinking a classic tale of mimicry. Sci. 
News 139:348-349. 


Received for publication 20 June 1992; revised and accepted 7 February 1994. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 199-204 


FIELD OBSERVATIONS OF MATINGS BETWEEN FEMALE 
LIMENITIS ARCHIPPUS AND MALE L. ARTHEMIS 
SUBSPECIES (NYMPHALIDAE) 


CHARLES V. COVELL, JR. 
Department of Biology, University of Louisville, Kentucky 40292, USA 


Abstract. Natural matings of female viceroy butterflies, Limenitis archippus, with 
males of L. arthemis astyanax in Kentucky and L. arthemis arthemis in Wisconsin are 
described and illustrated. These are the first field reports of female L. archippus paired 
with male L. arthemis. The interspecific pairing observed in Kentucky, plus the collection 
of two hybrid male “rubidus” Strecker at the same site in September 1980 and 1993 
suggest a high level of hybridization at that site. This seemingly high frequency is discussed 
in light of recently advanced hypotheses. 


Additional key words: Limenitis arthemis arthemis, L. arthemis astyanax, hybrid- 
ization. 


Natural hybridization among North American butterflies is appar- 
‘ently uncommon, except between Colias eurytheme and C. philodice 
(Silberglied 1973). However, hybridization recently has been observed 
with some frequency in the highly mimetic admiral butterflies Lime- 
nitis archippus (Cramer) and L. arthemis (Drury). The viceroy (L. 
archippus) has evolved appearance and behavior modeled after the 
monarch, Danaus plexippus (L.), and in Florida, the queen, D. gilippus 
(Cramer) (Ritland 1990). Limenitis arthemis astyanax is a Batesian 
mimic of the pipevine swallowtail, Battus philenor (L.) (Papilionidae), 
within the range of that swallowtail, grading northward to the non- 
mimetic, white-banded subspecies L. arthemis arthemis. Mimicry in 
these two eastern Limenitis species has been studied extensively in the 
laboratory (Platt & Brower 1968, Platt 1983). Platt et al. (1978) reported 
records of 10 wild-caught hybrids resulting from L. archippus x L. 
arthemis arthemis, known as “arthechippus’” Scudder, and 24 “rubi- 
dus” Strecker—the F, hybrid of crosses between L. archippus and L. 
arthemis astyanax. Platt (1987) raised the number of both hybrids to 
32, and Platt and Maudsley (1994) increase the number to 45. All known 
wild-caught hybrids are males. 


Collections and Observations in Kentucky and Wisconsin 


One of the records listed by Platt et al. (1978) was from Jefferson 
County, Kentucky, collected by R. Steilberg and J. Smith in September 
1948. That specimen is apparently lost. Three additional specimens 
have been taken more recently in Kentucky, and are in the University 
of Louisville collection. The late Siegfried Scholz collected one of these 
at Waverly Park near Valley Station, Jefferson County, on 21 October 
1978. I took another on the banks of the Mississippi River at Hickman, 


JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


200 


VOLUME 48, NUMBER 3 201 


Fulton County, Kentucky, on 14 September 1980. On 7 October 1989, 
at 1725 h, I encountered a nearly fresh female L. archippus mating 
with a ragged male L. arthemis astyanax (Fig. 1). This occurrence was 
only about 300 m south of the 1980 capture location. Robert V. Gregg 
also took a hybrid at the exact same site on 11 September 1993 (Figs. 
3, 4). 

Temperatures on the September capture dates in 1980 and 1993 were 
moderately high, and many pierids and other butterflies were recorded 
on those days. The day of the mating event in 1989 was cool, reaching 
a high of only about 17 degrees Celsius. Several other female viceroys 
were on bushes and trees nearby in basking postures. While not quan- 
tified, there were significantly more L. archippus than L. arthemis 
astyanax at the site. Abundant willow (Salix sp.; Salicaceae) was growing 
along the Mississippi River bank there, but I did not notice any Prunus 
species (Rosaceae). Very few butterflies of other species were still active. 

The mating pair was photographed by W. R. Black, Jr., and we 
observed the phenomenon until 1740 h when I decided to capture the 
paired insects. As soon as I approached them with a net, the female 
flew upward, carrying her mate with her. After attaining a height of 
about 45 m, she descended into a small tree, barely within reach of my 
net. After capturing them I placed the pair in a Ziploc bag where they 
continued mating until dark, an hour later. I took them back to Lou- 
isville, but found them moribund next morning. The female could not 
oviposit because a dried fluid (presumably hemolymph) had covering 
the posterior part of her abdomen. I dissected out several of the eggs 
but none hatched. 

In personal correspondence about this occurrence with A. P. Platt, I 
learned that this was the first field observation of a female L. archippus 
mating with a male L. arthemis astyanax. Later Platt informed me of 
a female viceroy seen mating with a male L. arthemis arthemis in Rusk 
County, Wisconsin (Fig. 2). In this case, photographs were taken by 
Steven Mertins of Ixonia, Wisconsin, after the discovery of the pair by 
his son Jake. The Mertins informed Platt that the mating was observed 
in “mid-afternoon” at the “end of August 1988.’ They did not collect 
the butterflies. This occurrence is the earliest record of a female L. 
archippus involved in interspecific mating. Platt et al. (1978) report 


—_— 


Fics. 1-4. 1, Female L. archippus mating with male L. arthemis astyanax, Hickman, 
Fulton County, Kentucky, 7 October 1989; 2, Female L. archippus mating with male L. 
arthemis arthemis, Rusk County, Wisconsin, August 1988; 3, Upperside of L. archippus 
x L. arthemis astyanax (“rubidus’’), Hickman, Fulton County, Kentucky, 11 September 
1993; 4, Underside of specimen shown in 3. 


202 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


three captures in Wisconsin of hybrid male ‘“rubidus” with features 
similar to Kentucky specimens (Figs. 3, 4). 


DISCUSSION 


The apparent frequency of hybridization in Limenitis at the western 
Kentucky site may lend support to hypotheses proposed by Ritland 
(1990). He pointed out that while records of hybrid admirals are rare 
over most of their sympatric ranges, there were 7 “rubidus’” recorded 
(2 seen or captured, and 5 reared from captured larvae) from a zone 
in southern Georgia to northern Florida during a 13-month period. 
Ritland also observed matings in nature between female L. arthemis 
astyanax and male L. archippus there. He indicates that these obser- 
vations “provide evidence of mate-choice breakdown’’—i.e., careless 
mate selection by L. arthemis astyanax females and consequent hy- 
bridization. This phenomenon may be more common in the southern 
Georgia/northern Florida area than elsewhere in their mutual ranges, 
as supported by additional evidence presented by Platt and Maudsley 
(1994). Ritland gives possible explanations as (1) more frequent matings 
between these species there than elsewhere, and (2) mating frequency 
not greater there, but F, hybrids “more viable than those elsewhere.” 
Segregation of the species by habitat determined by different foodplants 
is mentioned as a means of pre-mating segregation. Ritland points out 
that in his study area, there is little wild cherry—the usual foodplant 
of the red-spotted purple—and that the larvae feed on willows alongside 
L.archippus. Platt and D. Flaim have observed the larvae of L. arthemis 
subspecies feeding on Salicaceae in Maryland and New England as well. 
Midsummer senescence of wild cherry leaves could lead to increased 
oviposition on willows by L. arthemis astyanax by midsummer females 
producing the southern fall brood. This explanation would not hold for 
the hybrids reported by Ritland (1990) and Platt and Maudsely (1994) 
that eclosed prior to August. 

A third possible explanation is that the Georgia/Florida area rep- 
resents an intergrade zone between the northern orange L. archippus 
archippus and the dark brownish Florida viceroy, L. archippus flori- 
densis (Strecker) that mimics the queen, Danaus gilippus Cramer, 
instead of the monarch. In that area, normal mate selection behavior 
could be less discriminating, and L. arthemis astyanax may mate more 
readily with L. archippus. 

Against this background, the observation of one interspecific mating 
and two specimens of “‘rubidus’’ within the space of about five riverine 
acres in western Kentucky seems to lend support to the following con- 
ditions mentioned by Ritland as characteristic of a “hybridization hot 
spot.” 


VOLUME 48, NUMBER 3 203 


(1) Abundant willow but no black cherry were observed at the site 
(although black cherry could exist in some nearby woodland, subject 
to flooding as occurred in the summer of 1998). 

(2) Many more L. archippus than L. arthemis astyanax were seen 
at the site on each visit in September or October. The viceroy outnum- 
bered the red-spotted purple on 11 September 1998 by a ratio of 10:1 
(W. R. Black, Jr. pers. comm.). 

The first observation seems to qualify as what Ritland (pers. comm.) 
calls the “unusual habitat explanation.” With respect to probable use 
of willow as a larval foodplant by both species, the Fulton County, 
Kentucky site seems to match parts of the Georgia/Florida hybridiza- 
tion zone. Perhaps L. arthemis astyanax that mate successfully with L. 
archippus in the Kentucky site fed as larvae on willow rather than wild 
cherry. If so, pheromones of the two species might be more similar and 
isolating mechanisms diminished than if L. arthemis astyanax fed on 
wild cherry. In September and October very few L. arthemis astyanax 
were seen at that site, while the viceroy was common. Male L. arthemis 
astyanax might court female L. archippus in the absence of sufficient 
females of their own species. 

Less applicable to the Kentucky case is Ritland’s hypothesis that a 
higher rate of interspecific interaction might occur where viceroy wing 
color is unstable, as in the Georgia/Florida sites described by Ritland 
and by Platt and Maudsley (1994). Platt indicates viceroys with the 
contrastingly dark forewings can be found as far north as the Great 
Dismal Swamp on Virginia’s coastal plain. I have found very few in- 
dividuals with forewings darker than hindwings in Fulton County, 
Kentucky, and none with red spots on the underside as mentioned by 
Platt and Maudsley (1994). 

I cannot affirm that the frequency of hybridization between the two 
admiral species in Fulton County, Kentucky, approaches that reported 
from the Georgia/Florida habitat. However, three observations of such 
activity during a total of less than 12 hours of observation (1980-1993) 
over several seasons seems to indicate rather high frequency there. 

Platt et al. (1978) mention the prevalence of hybrid admirals occur- 
ring in late summer to early fall—toward the end of the flight season 
northward. The Kentucky observations and collections support that 
observation. 

My observations in Kentucky lead me to agree with Ritland (1990: 
171): “These arguments remain speculative because mate choice in 
viceroys and red-spotted purples is poorly understood; however, the 
proposed mechanisms identify several avenues of research that should 
be pursued in attempting to explain the elevated hybridization in this 
area.” The definition of hybridization “hot spot’’ should emerge as more 
field data are collected and assimilated. 


204 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


ACKNOWLEDGMENTS 


I thank A. P. Platt for providing the Wisconsin data and photos, and for his helpful 
comments. Thanks are due also to Jake and Steve Mertins for providing their color 
photograph from Wisconsin; to R. V. Gregg for donating his “rubidus” specimen to the 
University of Louisville Insect Collection; and to W. R. Black, Jr., and Scouts of Troop 
1, Paducah, Kentucky, for their assistance. My thanks also to L. P. Brower and D. Ritland 
for comments on the manuscript. 


LITERATURE CITED 


PLATT, A. P. 1983. Evolution of North America admiral butterflies. Bull. Entomol. Soc. 
Am. 29:10-22. 

. 1987. Recent observations of North American admirals (Lepidoptera: Nym- 
phalidae). Maryland Entomol. 3:18-20. 

PLATT, A. P. & L. P. BROWER. 1968. Mimetic versus disruptive coloration in intergrading 
populations of Limenitis arthemis and astyanax butterflies. Evolution 22:699-718. 

PLATT, A. P. & J. R. MAUDSLEY. 1994. Continued interspecific hybridization between 
Limenitis (Basilarchia) arthemis astyanax and L. archippus in the southeastern U.S. 
(Nymphalidae). J. Lepid. Soc. 48:190-198. 

PLATT, A. P., G. W. RAWSON & G. BALOGH. 1978. Interspecific hybridization between 
Limenitis arthemis astyanax and L. archippus (Nymphalidae). J. Lepid. Soc. 32: 
289-308. 

SILBERGLIED, R. E. 1973. Ultraviolet differences between the sulfur butterflies, Colias 
eurytheme and C. philodice, and a possible isolating mechanism. Nature 241:406— 
408. 

RITLAND, D. B. 1990. Localized interspecific hybridization between mimetic Limenitis 
butterflies (Nymphalidae) in Florida. J. Lepid. Soc. 44:163-173. 


Received for publication 25 October 1998; revised and accepted 19 April 1994. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 205-227 


A TECHNIQUE FOR SETTING AND MOUNTING 
MICROLEPIDOPTERA 


JEAN-FRANCOIS LANDRY AND BERNARD LANDRY 


Agriculture Canada, Centre for Land and Biological Resources Research, 
Central Experimental Farm, Ottawa, Ontario K1A 0C6, Canada 


ABSTRACT. Freshly collected and ammonia-killed microlepidoptera, pinned on min- 
utens, are spread in small, shallow, plastazote-lined boxes with grooves, using either small 
card points mounted on short pins or translucent setting paper strips to hold the wings. 
The method produces high-quality specimens, is fast, and uses compact, light-weight, 
inexpensive equipment. The method is also versatile in that any desired quality of setting, 
from preliminary, partial setting to the finest setting, can be attained with the same 
equipment with equal efficiency under any condition, whether at home or on collecting 
expeditions. The main steps of the method are illustrated. A technique for staging minuten- 
pinned specimens is also presented. 


Additional key words: spreading box, staging, double-mount, ammonia. 


During the past 90 years, several papers have presented, with various 
amounts of detail, techniques for preparing (pinning, setting, and 
mounting) microlepidoptera (e.g., Kearfott 1904, Calmbach 1921, 
Lhomme 1926, 1927a, 1927b, Amsel 1935, Holland 1937, Janse 1939, 
Janmoulle 1943, Charlson 1945, Lindquist 1956, Hodges 1958, Lewis 
1965, Tagestad 1974, Zimmerman 1978, Sokoloff 1980, Mikkola 1986). 
However, our contacts with many lepidopterists indicate that, at least 
in North America, good and simple techniques for preparing micro- 
lepidoptera are not well known. In fact, many North American lepi- 
dopterists do not even collect microlepidoptera as routinely as other 
Lepidoptera, in part because of the perceived inconvenience of pre- 
paring them. Microlepidoptera that are collected are often only the 
larger specimens, in groups such as pyraloids, tortricoids, and large 
gelechioids. 

The paucity of good quality microlepidoptera from North America 
in many collections is one of the causes for the very slow progress in 
systematic studies of the Nearctic fauna. Our knowledge of the tax- 
onomy and faunistics of many families of microlepidoptera is shockingly 
poor. A plea recently has been made for North American lepidopterists 
to take on the collection and study of microlepidoptera (De Benedictis 
1993). Of course the first step in this endeavour is to acquire a good 
and efficient technique for preparing specimens. 

There are probably nearly as many ways of preparing microlepi- 
doptera as there are individuals collecting them. The basic method of 
spreading microlepidoptera is the same as for larger Lepidoptera. How- 


206 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


ever, some adjustments, both at the time of collecting and of prepa- 
ration, and in the equipment, are needed because of the small size and 
fragility of microlepidoptera. 

Over the years we have tried every different method and variation 
of preparation for microlepidoptera that we came to know. While most 
techniques can yield high-quality specimens, many suffer from being 
relatively slow or requiring somewhat cumbersome equipment (e.g., 
spreading boards) ill-suited for prolonged field work under difficult 
conditions. We sought to develop a technique that offsets these prob- 
lems, i.e. one that is rapid and usable under any condition with equal 
efficiency, yet versatile with respect to the quality of preparation desired 
by the collector. An earlier version of the technique described here was 
published in French by Landry (1991) but we have modified it slightly, 
with some additions. 

Our method actually combines elements from other methods em- 
ployed by microlepidopterists, with added refinements. It is based on 
the concept of setting microlepidoptera on the bottom of a box, which 
can be traced back at least to Amsel (1935). Modern materials, especially 
dense polyethylene foam, dramatically enhance the results of Amsel’s 
method. Partial spreading in such boxes is now used by many micro- 
lepidopterists on collecting trips (Zimmerman 1978:50—-59, Nielsen 1980). 
The main shortcoming of partial spreading is that special specimens, 
such as types of new species or those needed for photography, may 
need subsequent relaxation for final spreading. The technique exposed 
here offers the possibility of a full range of quality of preparations, 
from unspread to fully spread with as much care as a perfectionist may 
wish, all with the same equipment and with hardly any extra time. The 
technique may be used in the field, in the lab, or at home. The necessary 
equipment is very compact, light-weight, inexpensive, and easily made. 
We have tested the method with tens of thousands of microlepidoptera 
over the past few years, under conditions varying from local day trips 
to month-long expeditions in the tropics (including camping). 

In addition to the actual technique of setting microlepidoptera, we 
offer some suggestions for handling specimens when they are collected 
in the field, and for staging (double-mounting) spread specimens. Ap- 
propriate handling of collected microlepidoptera is as critical as the 
actual setting in obtaining high quality specimens, and so is the final 
staging to insure safe preservation in subsequent handling. 


COLLECTING 


The facility and rapidity of the technique outlined here rests on 
working with the freshest specimens possible. Moths are placed indi- 


VOLUME 48, NUMBER 3 207 


vidually in glass vials upon collecting and kept alive until the time of 
pinning and setting. Upon returning from the field, vials are stored in 
a cool, dark place if the specimens cannot be prepared immediately. 
The ideal place is the refrigerator, or a cooler box if one is on a prolonged 
field trip. We have been able to keep specimens alive for up to five 
days in this manner, although we recommend delaying as little as 
possible (some moths will begin to show some wear even after 1-2 days 
in the refrigerator). Refrigeration is particularly useful if one has had 
a large catch on one day and there is not enough time to prepare all 
specimens immediately after they have been collected. We recommend 
preparing the smallest microlepidoptera as soon as possible, as they will 
die more quickly from dehydration. Once dead, small moths tend to 
dry very quickly and become difficult to relax and spread. In the humid 
tropics, small microlepidoptera will dehydrate quickly inside vials (of- 
ten in just a few hours) and are best set as soon as possible. Always 
begin by preparing the smallest specimens first, working up to larger 
ones. Refrigeration, even if available, should probably not be used for 
tropical microlepidoptera from those regions that seldom experience 
temperatures below 10°C, because the relative cold will kill many of 
them. 

Vials. Collecting vials should preferably be made of glass and close 
with an easily removable stopper that can be opened with a single hand 
(the other may be busy holding a net). We use glass vials that are 65 
mm long and 19 mm in diameter, closed with a rubber stopper. Stoppers 
should be as little wedge-shaped as possible, otherwise smaller micro- 
lepidoptera will crawl in the space between the stopper and the vial 
neck and damage themselves. We carry about 100 vials for most day- 
time collecting, at least twice as many for night-time collecting at a 
light. Experience will dictate the adequate supply. During day-time 
collecting, care must be taken that the vials are not exposed directly to 
or heated by sunlight, otherwise the moths will quickly die and dry. If 
possible, avoid plastic vials (snap-cap type), especially with the smaller 
specimens, because the static charge that such vials accumulate through 
handling and friction will damage the squamous cover of the moths 
and increase the rate of wear. 

A word of caution is necessary if one is setting reared specimens: 
never set a freshly emerged moth. Allow at least 24 hours (longer if a 
genitalia dissection may be required) for the moth to harden sufficiently. 
Without this precaution, wings may curl, crumple, or droop after re- 
moval from the setting box, and if the genitalia are later dissected, 
structures will be insufficiently sclerotized and difficult to prepare ad- 
equately. 


208 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


POISON AND KILLING TUBES 


The choice of poison is, of course, a matter of personal preference, 
availability, etc. We strongly reeommend ammonia (ammonium hy- 
droxide): it has a quick knock-down action and leaves freshly killed 
specimens beautifully relaxed and ready to be spread immediately with 
the greatest ease. We have tried other killing agents and methods, but 
ammonia is the one that has given us the best results. The ammonium 
hydroxide solution should be as concentrated as possible. A laboratory- 
grade solution containing about 30% ammonia and 70% water is pref- 
erable because it has a very fast knock-down action. Household am- 
monia, generally a murky liquid, is weaker and unsuitable. 

For killing tubes, we use glass tubes closed with cork stoppers into 
which a small microvial is inserted, loosely stuffed with cotton (Fig. 1). 
Five to ten minutes before using a tube, the cotton is imbibed with a 
few drops of ammonia solution, and the tube closed to let the ammonia 
concentration rise. This type of killing tube offers nothing inside against 
which struggling moths may rub; the disadvantage is that the tubes 
need to be recharged more frequently, approximately once every 2-3 
hours of continuous use (when opened several times periodically). When 
setting large numbers of microlepidoptera, we use up to 10 tubes at a 
time to minimize recharging, and place only 2-3 moths per tube at a 
time. It is essential to check for and wipe traces of moisture or sweating 
on the walls of the killing tubes. Charged tubes may be laid on their 
side to prevent any ammonia from possibly running down the sides, 
although this will not be a problem if a modest quantity is used. When 
tubes are not in use for more than a day or so, it is preterable to leave 
them open and remove the cotton swab from the stopper to allow then 
to dry thoroughly. 

Ammonia has a few disadvantages: it tends to sweat in a tube if an 
excessive quantity is used or if it is too warm (tubes must not be exposed 
to heat or direct sunlight)—but this is a disadvantage common to most 
liquid poisons; it loses strength relatively rapidly in a frequently open 
tube; and fumes are choking, irritating. Weak ammonia must not be 
used for moths with green, red, or orange pigments because the long 
exposure needed to kill them may cause discoloration. If the ammonia 
is strong though, this is not a problem providing that the moths are 
removed as soon as they are dead. In case of doubt about possible 
discoloration, one should use another poison, preferably ethyl acetate 
(subsequent relaxation may be necessary). With strong, concentrated 
ammonia, we have not had discoloration problems. Generally we have 
found that the advantages of ammonia far outweighed its disadvantages, 
none of which presented a real problem if it was used with the pre- 


VOLUME 48, NUMBER 3 209 


cork 


micro vial 
razor blade metal ruler 


or strip 


absorbant 
cotton 


spacer board 


Fics. 1-5. Materials required for preparing microlepidoptera. 1, Killing tube; 2, 
superimposed, offset rulers to cut symmetrical V-shaped grooves, A = B for 45° grooves; 
3, curved forceps used to handle minutens; 4, bent standard pin used to assist pinning 


and spreading; 5, card triangle mounted on shortened pin used to hold set wing in the 
point method. 


cautions outlined above, and that it was no more inconvenient to use 
than any other poison. 

Recently we have experimented with a solid form of ammonia, am- 
monium carbonate, a salt with the appearance of cyanide crystals. Upon 


210 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


contact with the ambient humidity, the crystals decompose into gaseous 
ammonia, carbon dioxide and water vapor (Gilligan and Gilligan 1990). 
Killing tubes are made simply by packing a 1-2 cm thick layer of 
crystals in the bottom and covering them with a smooth, porous material 
[e.g. artificial foam sponge (Gilligan and Gilligan 1990)]. We used plastic 
caps (from snap-cap vials) punctured with many minute pin holes to 
cover the crystals. Plaster cannot be used because the water it contains 
will instantly dissolve all the crystals and produce all the ammonia at 
once. We obtained satisfactory results with ammonium carbonate if 
used for small numbers of specimens. Disadvantages are that the ra- 
pidity of killing decreases markedly compared to liquid ammonia if 
one opens the tubes frequently; also if there are too many specimens 
in a tube and it is warm, the moisture content may rise to the point 
where, upon cooling, crystals may form on the specimens; such crystals 
are then very difficult to remove. For these reasons we find ammonium 
carbonate less satisfactory than ammonium hydroxide. 

Ethyl acetate also works well but we found that it has a tendency to 
stiffen many microlepidoptera if they are left in the killing tube a few 
minutes too long; hence, some relaxation is sometimes necessary. Like 
ammonia, it is volatile, and tubes need frequent recharging and may 
“sweat” if heated. It is also flammable and will dissolve some plastics. 
Generally, we have found ethyl acetate to be less satisfactory than 
ammonia in quickly producing ready-to-spread specimens. 


KILLING 


Remove the cork, insert one moth, close the cork. Repeat with other 
tubes. When there is one moth in each tube, start again with the first 
tube, ensuring that the moth is stunned. Continue until there are 2-3 
moths per tube. Stunning takes less than five seconds when the ammonia 
is strong but may stretch to 10-15 seconds after tubes have been opened 
several times. Moths should be left in the killing tubes for at least 15 
minutes to ensure they are dead. Very small moths (Nepticulidae, small 
Gracillariidae, for example) can be removed sooner. A time saving 
strategy in the subsequent setting operations is to segregate specimens 
by size at the killing stage. This way, at the setting stage, one does not 
have to switch back and forth among various spreading boxes with 
different groove widths. 


SETTING EQUIPMENT 


Spreading boxes (Figs. 6, 17-18). We use shallow, clear polystyrene 
plastic boxes; currently we have two sizes, llem x llem X 2cm, and 


VOLUME 48, NUMBER 3 . | 211 


— 


Fic. 6. Spreading box. The actual lid is used as bottom on which the plastazote is 
glued. Scale in cm. 


12cm X 8cm X 2cm, obtained from different suppliers. Actual dimen- 
sions are not important, as long as boxes are relatively small, preferably 
shallow (for compactness), with a low-edge lid, rigid, and relatively air- 
tight (or pest-proof). 

For a spreading surface we use Plastazote®, a dense, smooth poly- 
ethylene foam. We found this material best because it affords the 
following advantages: the surface acquires a small static charge through 
handling, which helps wings cling slightly and facilitates spreading; it 
grips the pins firmly and leaves no pin holes; it sustains hardly any 
wear. | 

A 1-cm thick piece of plastazote is glued inside the lid of a spreading 
box (we use all-purpose, non-toxic white glue). Gluing the foam inside 
the lid (using the bottom as lid) eliminates edges to the spreading 
surface, greatly facilitates work of the hands, and maximizes use of the 
spreading surface. 

Before gluing the foam into the boxes, we cut three or four V-shaped 
grooves with a razor blade. To obtain grooves with perfectly symmet- 
rical sides, we use two metal rulers or strips, with one being taped on 
top of the other and propped up by a thin board; the edge of the top 
ruler is offset from the edge of the lower one by a distance equal to 
that of the ruler + board thickness (Fig. 2). To cut, the blade is slanted 


PA JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


~ 6-8mm 


Fics. 7-8. 7, Inserting the minuten while holding the body with a bent standard pin; 
8, Minuten-pinned specimen, showing approximate height on 1-cm long minuten. 


and abutts both edges. Symmetrical grooves facilitate spreading. We 
use a series of spreading boxes with various groove widths, these varying 
from 1-5 mm (2mm and 8mm are the most frequently used widths). 
It is not necessary to have square grooves with vertical sides, as on 
standard spreading boards. In fact, the sides of V-shaped grooves often 
provide direct support for the abdomen. 

Minuten pins. Use of minuten pins involves subsequent staging or 


VOLUME 48, NUMBER 3 . 213 


double-mounting, so this is distasteful to many a lepidopterist. Whatever 
the perceived difficulty, inconvenience, or time factor involved, we 
emphasize that this is by far the best and safest way of obtaining fine- 
quality microlepidoptera. Double-mounted specimens can sustain 
rougher handling without damage and are far less likely to lose their 
abdomen, a very frequent problem with microlepidoptera that are 
mounted on fine standard pins (00 or 000), which are very springy. The 
genitalia are critical for the specific determination of numerous species 
of microlepidoptera, hence the abdomen must not be lost. 

There are different qualities of minutens available on the market. 
For the best results, and to avoid frustration, one should use the best 
quality stainless steel minutens. Avoid black-enameled minutens, which 
have a tendency to rust (guaranteed if one is in the tropics) and have 
tips that more easily “hook” (being made of softer metal). The differ- 
ence in price between stainless steel and black-enameled minutens is 
small. Diameters of the most useful sizes are 0.20 mm, 0.15 mm, and 
more rarely 0.10 mm (for nepticulids and other tiny microlepidoptera); 
some British brands label their minutens Al (0.14 mm) and B1 (0.19 
mm) (1 referring to the shortest length, usually 10-12 mm). 

Most minutens are excessively long and must be shortened down to 
no more than about | cm for the larger ones (0.20 mm) or 6-7 mm for 
the finer ones (0.15 mm and 0.10 mm). If minutens are not shortened, 
the excess length jutting either above or below the specimens will greatly 
increase the risk of breakage or damage during handling of the double- 
mounts (fingers pinching the minuten while grasping the stage-sup- 
porting pin will spring the specimen and likely send parts flying, most 
commonly the weakly-attached, all-precious abdomen). A rapid method 
of shortening a large number of minutens is to cut narrow strips of 
plastazote (often the latter’s thickness is conveniently 1 cm or 7 mm), 
to insert minutens all the way through the strips (ensuring that their 
tips do not extrude), and trimming off the excess length close to the 
strip surface with good scissors or pin cutters. To maximize efficiency 
later in the setting process, we prepare large quantities of trimmed 
minutens in advance. Minuten-loaded plastazote strips can be packed 
side by side in an insect mounting tray or small shallow cardboard box. 
A protective layer of plastazote is glued on the bottom of the tray or 
box. Strips are then laid upright, side by side, and held in place with 
pins inserted through the sides of the tray or box; any remaining space 
can be filled with plastazote. Use a box narrow enough for the holding 
pins to pierce through at least half of the strips from one side. 

Tools. We use curved forceps for handling minuten pins (Fig. 3). 
The inner surface of the grasping end must be smooth (not striate). 
While fine straight forceps could be used, we found curved forceps to 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


214 


WSS 


SS=s 


SS 


AN 


VOLUME 48, NUMBER 3 ; 215 


be a much more versatile tool for the task. A large standard pin (e.g. 
no. 4) bent at an obtuse angle (Fig. 4) provides an inexpensive tool, 
instead of a second pair of forceps, to help in holding the specimens 
or the wings during pinning and setting. It is important to use a pin 
that is not too fine because the point may catch and rip into the wings 
too easily. 

Setting triangles (Fig. 5). Triangles are used to hold the wings in 
place once they are spread. They are made with a point punch (of the 
type commonly used for mounting small insects) from moderately thick, 
very smooth or glossy card and inserted no more than half way up on 
short pins. We use two sizes of triangles [7 mm long with pointed end 
(Fig. 17) and 10 mm long with truncate end (Figs. 10—12)] for different 
sizes of microlepidoptera. We mount them on pins no. 00 cut down to 
1 cm in length (trimmed the same way as the minutens). Do not use 
minutens for mounting triangles because they are too fine to insert 
easily into the relatively thick card stock of the triangles. When mount- 
ing triangles, check that the side with rough edges (produced by the 
punch on the underside of the paper) is turned upwards (check with a 
magnifying lens if necessary). If this simple precaution is not taken, 
much damage to the wing scales will occur because of the rough edges 
of the triangles. As for minutens, a large supply of mounted triangles 
should be readied, pinned in shallow boxes. Triangles are re-used in- 
definitely or until they become loose on the supporting pins. 

Pinning pad. White cotton fabric folded several times into a pad 
about 1 cm thick and 10 cm x 10 cm makes an ideal surface to pin 
microlepidoptera. The fabric must be as soft as possible. This surface 
prevents specimen compression when pinning and the fabric fibres tend 
to hook the moth claws, thus reducing slippage. Avoid paper towels of 
any kind, they are usually too rough. The thickness of the pad must be 
greater than the length of the minutens so that the pad does not have 
to be lifted up when pushing the minutens through the specimens. 

Humid container. This serves to hold pinned specimens to prevent 
them from drying while they await setting. If this precaution is not 
taken, the smaller microlepidoptera will begin to dry in a mere few 


— 


Fics. 9-12. Setting with the point method. 9, Inserting the specimen into the groove 
with the wings partly opened; 10, Moving one set of wings forward with the point- 
holding pin; note the antenna held in position with a minuten, 11, Setting the wing into 
position with the mounted point while holding it with the bent pin; 12, Repeating the 
operation with the other side. 


216 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


LZ 
oy 
fi. 


VOLUME 48, NUMBER 3 . 217 


minutes in dry air and become difficult to spread by the time one gets 
to the last few of a batch. The container is simply made from a plastic 
petri dish or similar small plastic dish or box with a loosely fitting lid. 
The bottom of the dish is lined with wetted tissue or filter paper. A 
small plastazote pad serves to hold specimens. In very dry conditions, 
the inside of the lid may be lightly misted to increase ambient moisture 
in the container (too much moisture could drip on the specimens). 


PINNING AND SETTING 


For best results (and less eye strain) the pinning and setting operations 
should be done with magnifying lenses or under a low-power stereo- 
scope (up to about 5x). 

Pour the freshly killed moths on the cotton pad and pin them. Insert 
the minuten through the center of the mesothorax (mesoscutum) or at 
the suture between the mesoscutum and mesoscutellum (the mesos- 
cutellum is the roughly triangular or diamond-shaped area behind the 
center of the mesothorax). Try to keep the pin in line with the center 
of the thorax, otherwise the wing muscles may become transfixed, which 
renders spreading more difficult. To ensure that a specimen is squarely 
pinned, apply very slight pressure on its dorsum with the tip of the 
bent pin (or another curved forceps) to prevent the body from rolling 
sideways while the minuten is inserted into the mesothorax (Fig. 7). 
The minuten must be inserted far down so as to leave no more than 
about 2 mm protruding above the moth, enough to manipulate it com- 
fortably with forceps (Fig. 8). Of course, the height of specimens with 
unusual structural modifications such as long palpi recurved over the 
body or thoracic crests should be adjusted appropriately in order to 
leave sufficient minuten length for the forceps; such specimens may 
require longer (untrimmed) minutens. 

Place pinned specimens in the humid container. Prior to this, if one 
wishes, the wings may be partly opened by gently blowing on them 
from behind the moth with a slight puff of breath. Before proceeding 
with setting, another series of specimens is transferred to the killing 
tubes. Hence, there will be specimens ready for pinning when the first 
batch has been set. We usually proceed in batches of no more than 15- 
25 moths. 


—_— 


Fics. 13-16. Setting with the paper method. 13, Moving one set of wings forward 
with a minuten while lifting the paper strip with the bent pin; note the minuten holding 
the antenna; 14, Combing the fringe; arrow indicates direction of combing movement; 
the combing minuten touches the tip of the fringe lightly; 15, Pinning the paper strip 
down to secure the wing into position; the minuten holding the antenna may be removed 
as it is usually no longer necessary; 16, Set specimen. 


218 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Take specimens out of the humid container singly for setting. If the 
wings are still closed, gently blow on them from behind, then insert 
the specimen into the groove (Fig. 9). Lift the wings and partly push 
them forward with the tip of the closed curved forceps inserted beneath 
the wings. Tuck the legs into the groove. With a minuten position the 
antennae so that they form a widely obtuse V, holding them temporarily 
by placing minutens behind their base. If the fringes are matted, lift 
the wings a little and comb the fringes by brushing them with the tip 
of the triangle’s pin in a movement going from the apex of the wings 
toward the body. 

To fix the wings into position, we use two different procedures. 

(1) Points method (Figs. 10-12). This method may be a bit faster 
than the paper method (see below). Although excellent, it sometimes 
gives slightly inferior results, and makes it more difficult to set the 
antennae properly. 

Using a mounted triangle, bring one pair of wings forward by pushing 
on the hind margin of the forewing with the tip of the pin. Usually, if 
this movement is delicately executed, both wings will move together 
because of the coupling. Do not pierce the wings. While holding the 
wings into position with a slight pressure of the bent pin held in the 
other hand, put the triangle on top of the wings as close to the apex of 
the hindwing as possible and push it down sufficiently to immobilise 
the wings. The triangle must lie flat against the wing surface and must 
not be pressed down too strongly or it will leave a mark. It may be 
necessary to adjust the position of the hindwing slightly, which some- 
times will be a little too far back or too far forward. One or more 
triangles may be added to better hold the wings of larger or broad- 
winged microlepidoptera or to prevent them from curling up. 

Repeat the procedure for the other side. To prevent set specimens 
from hindering hand work over the spreading box surface, it is best to 
proceed in transverse rows instead of filling one groove after another. 

For someone having difficulty using both hands simultaneously, the 
following variation may be applied: using a mounted triangle as outlined 
above, move a pair of wings only halfway forward then insert the 
triangle over the wings just sufficiently to prevent the wings from 
slipping back but ensuring that they can still be moved; with a minuten 
move the wings into their final position (the wings should stay in place) 
and with the forceps maintained closed, gently push down the top of 
the pin holding the triangle until the wings are flat. Positioning of the 
wings in this way may have to be done in several stages for some 
specimens. The other hand may hold the spreading box. With this 
variation, one can proceed by filling one groove after another if desired. 

(2) Paper method (Figs. 13—16). The second procedure uses small 


VOLUME 48, NUMBER 3 : | 219 


strips of thin, translucent setting paper and is essentially similar to the 
standard technique used to spread larger Lepidoptera on a normal 
setting board. The paper strips are held down with 0.20 minutens. This 
technique can yield the finest specimens because the entire surface of 
the wings is held flat, and the antennae can be set properly with ease. 
It is a little more cumbersome and may take a little more time depending 
on individual ability. In our own experience, however, it takes about 
the same amount of time as the triangle technique, if one has prepared 
and has ready the necessary materials, such as pre-cut pieces of setting 
paper and minutens. 

Cut many small pieces of setting paper, just long enough to cover 
the antennae and one set of wings, prior to spreading. For most mi- 
crolepidoptera, the strips we use are about 1-1.5 cm long and about 5 
mm wide. Four minutens are usually needed to spread one moth. For 
increased speed, sets of four paper-holding minutens may be pinned 
beside each groove of an entire row before the setting begins. When 
one row has been filled with specimens, another series of minutens is 
placed along the next row, and so on. 

After pinning the moth and having set the antennae as described 
above, pick up a paper strip by stabbing it with a minuten and pin it 
just ahead of the antenna to cover the half-opened wings. Check that 
the curvature of the paper faces upward. With the bent no. 4 pin (or 
another pair of curved forceps) held in one hand, slightly lift the pos- 
terior end of the paper from beneath. With another minuten held with 
curved forceps in the other hand and working from behind the hind 
margin of the forewing, push the wings into position. When both wings 
are positioned, drop the paper strip, hold it down with the tip of the 
bent no. 4 and pin it behind the hindwing with a second minuten. 
Repeat on the other side. 

Choosing the appropriate groove width will facilitate spreading. A 
groove too narrow will force the legs up and put pressure on the thorax, 
thus hindering wing movement. A groove too wide will result in either 
the specimen swinging on the pin when the wings are pushed on one 
side, or in an insufficient portion of the wing surface resting flat. 

Before placing the specimen into the groove, the plastazote surface 
may be gently rubbed with the tip of the closed forceps to create a 
charge of static electricity which will help in spreading the wings. This 
is not necessary, however, if one is using the paper strip method, and 
it is not recommended with very small microlepidoptera such as nep- 
ticulids because the charge will be too strong and may push the wings 
up vertically. 

With a fresh, fully relaxed moth and some practice, the whole op- 
eration of pinning and setting takes no more 380-60 seconds. With 


220 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


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set specimens (larval cases beside reared specimens). Scale in cm. 


VOLUME 48, NUMBER 3 221 


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Fic. 19. Example of packed spreading box in which previously set and dry specimens 
are overlapped like shingles to conserve space. This 11 x 11 cm box contains 166 specimens. 


practice, specimens can be set quite closely behind one another into 
the grooves to conserve space (Figs. 17-18). 

Stunning prior to spreading is sometimes used, instead of killing, 
when time is short (Sokoloff 1980). If specimens are only anesthetized 
(stunned) prior to spreading, it is necessary to pin a small cotton swab 
imbibed with ammonia into the spreading box and close it for about 
15-20 minutes to kill the moths. If the spreading box is made of poly- 
styrene-base plastic, avoid ethyl acetate because it will dissolve the 
plastic. We do not use the stunning method because we find it incon- 
venient, especially in the field. 

Label the specimens as usual and leave them in the spreading boxes 
in a dry place for at least two weeks, or preferably for as long as possible. 
If one does not provide enough time for the specimens to dry, the tips 
of some wings may curl up or droop. In humid regions, it is advisable 
to secure a few crystals of 4-chloro-m-cresol in the boxes to prevent 
molding. Once the moths are dry, full boxes should be sealed tightly 
with tape until ready for staging. 

When in the field for an extended time and spreading boxes are in 
short supply, or to reduce the number of boxes being transported, space 
can be saved by removing specimens from the grooves after drying 


222 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


and packing them somewhat like shingles (Fig. 19). Specimens are 
pinned slanted in transverse rows, with the left wings of a specimen 
partly overlapping the right wings of the preceding one. This allows 
for large quantities of specimens to be stored in little space. An entire 
collection of several thousands of microlepidoptera can be carried in 
this way in a handbag on a plane instead of being placed in regular 
baggage, thus maximizing the safety of specimens that may represent 
months of field work in a remote region. 

Some authors have recommended heat-drying because, supposedly, 
moths that have been heat-dried will never have drooped wings (Amsel 
1935). This is, however, a delicate and risky operation that must be 
done very carefully with very low heat (ca. no more than 40° C). We 
have tried drying on a few occasions and are rather weary of it. We 
have noticed that several microlepidoptera tend to become a little greasy 
when dried with heat (noticeable under magnification). Another prob- 
lem is that the plastazote of the spreading boxes may warp slightly from 
being heated. We think that it is preferable to see some wing drooping 
occur later in the collection than risk damaging specimens in heat- 
. drying. Wing drooping will be minimized or virtually eliminated if 
specimens are allowed to remain set in the spreading boxes for an 
extended period. 


STAGING 


To be placed in collections, dry minuten-pinned microlepidoptera 
must be mounted individually on small rectangular blocks, which are 
inserted on standard (# 38 or 4) insect pins. This is referred to as staging 
or double-mounting. Specimens should always be mounted singly on a 
block, complete with all necessary labels on the supporting pin, except 
perhaps in cases of mated pairs which may be staged together. It is 
very annoying to find two or more microlepidoptera belonging to dif- 
ferent but superficially similar species that have been staged together 
with a single label; such specimens have to be remounted separately 
and new labels produced. Multiple mounts also increase the risk of 
misassociation of subsequently made genitalia slides. 

Staging blocks. It is more efficient to prepare large quantities of 
blocks in advance. Traditionally, blocks have been cut from strips of 
polypore fungi (especially from birch bracket fungus). Normally it is 
easy to procure polypore strips from naturalist supply houses, but pe- 
riodically they tend to become very difficult to obtain. 

Plastazote provides a superior substitute. It is comparatively inex- 
pensive, available in practically infinite supply, extremely regular in 
density, practically unalterable, and pest proof (we once had a supply 
of polypore strips heavily infested with ciid beetles). Plastazote allows 


VOLUME 48, NUMBER 3 223 


the finest minutens to be inserted without effort and provides remark- 
able protection from shocks and vibrations. Other materials such as 
balsa, cork, and polystyrene-based foam (‘‘styrofoam’’) should be avoid- 
ed because they are either too hard to insert the minuten without risking 
damage or are not rubbery enough to hold firmly the pin and the 
minuten (the latter is a problem of balsa and polystyrene-based foams, 
on which minutens frequently become loose). Blocks made of a silicon 
rubber compound are used by some but their durability is uncertain 
in insect drawers where they may be affected by fumigants; we have 
seen a set of such blocks that were about 15 years old and that exuded 
a greasy substance which seeped up the minutens and coated the spec- 
imens. It is also harder to insert a minuten into silicon rubber, which 
is a springy material. 

The length of the blocks varies with the size of the specimens. Ideally, 
we think that they should be about as long as the length of the moth 
from its head to the tip of its abdomen plus about 3 mm to provide 
space for the legs and the supporting pin. The width and height vary 
little and are from 2-3 mm (width) and from 2-4 mm (height). We 
recommend the use of as few sizes of blocks as necessary to maintain 
some uniformity to the collection. A cutting board with preset guides 
and mounted razor blade can be made to speed the cutting of large 
numbers of uniformly sized blocks. It is essential to mount the blocks 
on standard pins prior to double-mounting the moths. Staging blocks 
must be inserted up to a height that will leave adequate clearance 
between the specimen and the head of the supporting pin to allow for 
safe handling of the whole mount (Fig. 22); we recommend at least a 
lcm clearance. | 

Staging procedure (Figs. 20-21). To facilitate staging, use one pair 
of forceps with curved tips and another with broad, flattened tips. 

With the flat-tip forceps, hold the pinned block in front of you. With 
the curved forceps, take the specimen by holding the minuten from 
beneath the specimen and insert slightly into the block. Check that the 
plane of the wings is perpendicular with the axis of the pin and adjust 
the inclination if necessary. Still grasping the minuten from beneath 
the specimen, pull it down into the block to the point where the venter 
of the moth is about 1 mm from the surface of the block. 

Holding the minuten from beneath the specimen for insertion is 
especially critical if one is using polypore blocks. Polypore blocks vary 
markedly in hardness and pushing the minuten down while grasping 
it from above the specimen may cause the minuten to bend or spring, 
usually resulting in damage to the moth. Using plastazote blocks gen- 
erally obviates this danger but grasping the minuten below the moth 
reduces the risk of damage in case of slippage of the forceps. 


224 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


212mm 


 3-5mm 


Fics. 20-22. Staging or double-mounting. 20, Holding with flat-tipped forceps a 
staging block mounted on a standard pin; 21, Inserting the specimen on the stage, clasping 
the minuten from below the specimen; 22, Staged specimen, showing good heights for 
safe subsequent handling. 


It is important to insert the minuten as far down as possible, while 
not touching the stage, in order to secure the specimen (Fig. 22). Spec- 
imens protruding high on the block risk getting damaged in subsequent 
handling as much as those with overly long minutens jutting high above 


the body. 


VOLUME 48, NUMBER 3 225 


FINAL REMARKS 


The techniques described above may seem laborious, but what takes 
many words to explain is actually executed in just a few seconds. With 
some practice, one can easily pin and set up to 30-40 microlepidoptera 
of fine quality per hour. 

If there is no time or desire to fully spread all the moths that are 
collected, one may at least spread the wings partly and brush the fringes. 
Provisional spreading (Amsel 1935, Zimmerman 1978: pp. 48-ff, Nielsen 
1980, Mikkola 1986), with subsequent relaxation and spreading if nec- 
essary or desired, is a good compromise where time is short such as 
during expeditions aiming at sampling as many specimens as possible. 
Damaged or rubbed specimens that may be worth collecting for some 
reason may be partially spread to save time. 

Generally we do not use light traps and prefer to collect microle- 
pidoptera at light on a sheet. Although light traps afford several ad- 
vantages in sampling and are often necessary for surveys, we find that 
one is easily overwhelmed by the abundance of specimens so obtained, 
that a significant amount of time is necessary to sort the microlepidop- 
tera from other Lepidoptera and insects, and that most specimens sus- 
tain a certain amount of rubbing and damage. If there is no time to 
relax and set trap-collected specimens right away, they should be placed 
on slightly damp cotton in tight containers and kept in a freezer. 

Methods that involve killing the specimens immediately upon cap- 
turing them (as in light traps) and storing them for an indeterminate 
period of time (e.g. by freezing), generally necessitate some period of 
relaxation in a humid chamber before proper setting can be performed. 
Such specimens are usually not quite as easy to spread as freshly killed 
specimens and are not ideal for the point-setting technique described 
above, although satisfactory results can be obtained with adequate re- 
laxation and using the paper-strip technique. Specimens that have dried 
unspread usually cannot be subsequently relaxed and spread. Some 
lepidopterists who have tried our technique complained that it was not 
quite as easy as we told them but, when pressed for details of how they 
proceeded, most conceded that they had killed their specimens upon 
collecting and spread them a little later without relaxation. We reiterate 
that working from fresh, live specimens killed just before setting is 
central to the ease and rapidity with which microlepidoptera can be 
set with the technique described here, and to obtaining high-quality 
specimens. Of course, some experience is necessary to achieve the best 
results; one is unlikely to obtain perfect microlepidoptera after at- 
tempting to set only a dozen specimens. 

It is a truism that fine, well-prepared specimens are easier to identify. 


226 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


This is particularly true for microlepidoptera, whose small size puts 
them at a disadvantage over the larger Lepidoptera when it comes to 
studying them (incidentally, lepidopterists facing space limitations to 
house their collection of macros should seriously consider taking up the 
collection of microlepidoptera!). Many well prepared microlepidoptera 
can be recognized at a glance. On the other hand, rubbed, damaged, 
or badly mounted specimens may be quite difficult to recognize, even 
to family, particularly if they are unspread. 

Unavoidably, processing microlepidoptera soon after their collecting 
will take more time and seem more laborious than for larger Lepidop- 
tera that are simply papered or pinned for subsequent setting. It can 
be argued, however, that the time involved strictly in spreading mi- 
crolepidoptera is no more than for spreading macros; in fact spreading 
microlepidoptera is faster. The main difference is that one should do 
it right at the time of collecting for best results. The resulting quality 
of the specimens makes it well worth the effort. 


ACKNOWLEDGMENTS 


Several lepidopterists made suggestions and comments on our technique and on spread- 
ing microlepidoptera in general, and/or have encouraged us over the years to publish 
our method. In particular, we are indebted to Vitor Becker, Bengt Bengtsson, Don Davis, 
John De Benedictis, Michael Fibiger, John Grehan, Ron Hodges, Ole Karsholt, Eric 
Metzler, Kauri Mikkola, John Morton, Jerry Powell, Tony Roberts, Klaus Sattler, Dave 
Wagner, Monty Wood, and Don Wright. We thank Eric Metzler for making us aware 
of ammonium carbonate, and Cees Gielis for testing it under harsh field conditions. We 
thank Dave Moorehouse for his assistance in preparing the figures. Ole Karsholt, Jeff 
Cumming, Mike Sharkey, Kevin Tuck, and an anonymous reviewer, provided many 
useful comments on the manuscript. 


LITERATURE CITED 


AMSEL, H.-G. 1935. Comment préparer les microlépidoptéres secs. Amat. Papillons 7: 
238-240. 

CALMBACH, V. 1921 [1923]. Die Praparation der Mikrolepidopteren, unter besonderer 
Beriicksichtigung der kleinsten Arten unter den Kleinen. Entomol. Zeits. 35: 35-36. 

CHARLSON, S. 1945. Setting Microlepidoptera. The Amateur Entomologists’ Society 
Leaflet no. 14. London, England. 4 pp. 

DE BENEDICTIS, J.A. 1993. Why not collect micros?: Getting started. News Lepid. Soc. 
1993(3): 69-70. 

GILLIGAN, T. & M. GILLIGAN. 1990. A new killing jar. Ohio Lepid. 12: 62. 

Hopces, R. W. 1958. A method for preparing fresh microlepidoptera for spreading. 
Lepid. News 12: 205. 

HOLLAND, W. J. 1937. The moth book. Doubleday, Doran & Co., New York. xxiv + 
479 pp. 

JANMOULLE, E. 1943. Récolte et préparation des Microlépidoptéres. Bull. Mens. Nat. 
Belg. 7: 1-6. 

JANSE, A. J. T. 1939. On collecting, preserving and packing lepidopterous insects. J. 
Entomol. Soc. South. Africa 2: 176-180. 
KEARFOTT, W. D. 1904. Micro-Lepidoptera—Suggestions. Entomol. News 15: 89-96. 
LANDRY, J.-F. 1991. Récolte et préparation des Microlépidoptéres. Fabreries 16: 1-21. 


VOLUME 48, NUMBER 3 | 227 


LEwIs, G. G. 1965. A new technique for spreading minute moths. J. Lepid. Soc. 19: 
115-116. 

LHOMME, L. 1926. Chasse, préparation et conservation des papillons de petite taille. 
Amat. Papillons 3: 149-158. 

1927a. Chasse, préparation et conservation des papillons de petite taille (suite). 

Amat. Papillons 3: 166-176. 

1927b. Chasse, préparation et conservation des papillons de petite taille (suite). 
Amat. Papillons 3: 181-191. 

LINDQUIST, O. H. 1956. A technique for pinning and spreading small microlepidoptera. 
Canad. Entomol. 88: 24-25. 

MIKKOLA, K. 1986. Tower-spreading, a handy method for provisional field-preparation 
of microlepidoptera. Not. Entomol. 66: 101-102. 

NIELSEN, E. S. 1980. Entomology. The Danish Scientific Expedition to Patagonia and 
Tierra del Fuego 1978-1979. Geogr. Tids. 80: 9-18. 

SOKOLOFF, P. 1980. Practical hints for collecting and studying the microlepidodptera. 
Amateur Entomol. 16: 1-40. 

TAGESTAD, A. D. 1974. A technique for mounting microlepidoptera. J. Kansas Entomol. 
Soc. 47: 26-30. 

ZIMMERMAN, E. C. 1978. Microlepidoptera. Insects of Hawaii, vol. 9. University Press 
of Hawaii, Honololu. xviii + 1903 pp. 


Received for publication 1 December 1998; revised and accepted 27 February 1994. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 228-247 


CHECKLIST OF SOUTH DAKOTA BUTTERFLIES 
(HESPERIOIDEA AND PAPILIONOIDEA) 


GARY M. MARRONE 
HCR-38, Box 4C, Fort Pierre, South Dakota 57532, USA 


ABSTRACT. This paper presents a list of 172 butterfly species (Hesperioidea and 
Papilionoidea) documented from South Dakota. The list was compiled from records of 
resident and non-resident collectors, published literature, and university collections. For 
each species all recorded counties and adult flight periods are given. 


Additional key words: distributions, Great Plains. 


This paper is the first comprehensive treatment of South Dakota’s 
butterfly fauna. An early list of butterfly species by Truman (1896, 
1897) covers an area near Volga, Brookings County. Many of Truman’s 
specimens are deposited in the H. C. Severin Insect Collection at South 
Dakota State University. Other regional works include those for Sieche 
Hollow State Park in Marshall and Roberts counties (Masters 1975), 
and the Black Hills (Conway 1966). Papers treating only a species or 
group of species include Ferris (1969, 1976, 1981, 1983), Grey et al. 
(1963), McCabe (1981), Nordin (1967, 1968), and Scott and Stanford 
(1981). Many records for the western half of the state, west of the 100th 
meridian, are mapped by Stanford (Ferris & Brown 1981). Stanford 
and Opler (1993) reported butterfly species occurrences (county dot 
maps) for the western United States, including all of South Dakota. 

South Dakota is known as the “land of infinite variety” because of 
the diversity of terrain, habitat, and climate. There is a general increase 
in elevation from 335 m in the southeast to 1035 m in the northwest; 
the highest point is Harney Peak (2209 m) in the Black Hills and the 
lowest is on the Minnesota River (293 m) in the northeast corner of the 
state. The state is mostly gently rolling grassland with the Missouri 
River dividing it from north to south into two nearly equal parts. The 
area east of the river is characterized by a glaciated topography with 
tall grass, low rolling hills, and many lakes and marshes. The western, 
semi-arid mixed grass prairie, portion of the state has deep valleys and 
broad upland flats and buttes. The Black Hills, on the extreme western 
edge of the state, may be regarded as an outpost of the Rocky Mountains. 
Mean annual precipitation ranges from 66 cm in the southeast and 
Black Hills, to less than 30.5 cm in the northwest. 

One hundred seventy-two species of butterflies are known to occur 
in South Dakota. Of these, 132 are residents and the remainder are 
immigrants or strays. Other species undoubtedly will be added to this 
list as collecting continues. The author encourages anyone who can 
provide additional information to contact him. 


229 


VOLUME 48, NUMBER 3 


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A “butterfly calendar,” illustrating adult flight periods, was developed 
based on 6,500 South Dakota butterfly records (Table 1). Data were 
lacking for some species so approximations were made. The number 
of species occurring per each month is as follows: 


Apr Ma un ul Au Sep _—_ Oct 
20 76 137 131 93 o3 17 


More than 2,500 county records, tabulated through December 1993, 
are presented. This equates to an average of 39 species from each of 
South Dakota’s 66 counties. County locations and names are shown in 
Fig. 1. The number of species known per county ranges from 11 for 
Edmunds County to 117 for Pennington County (Table 2). In general, 
the counties with the highest totals are those of the Black Hills which 
is a reflection of a greater sampling effort combined with a broad 
diversity of habitats. The counties with the fewest reported species are 
generally highly agricultural or have not been surveyed thoroughly. 
No species has been documented for all South Dakota counties; however, 
five species have been recorded for more than 60 of the 66 counties: 
Pieris rapae, Colias eurytheme, Colias philodice, Lycaeides melissa, 
and Cercyonis pegala. Twenty-seven counties have fewer than 80 spe- 
cies recorded. South Dakota’s total of 172 species ranks fourth among 
its bordering western Great Plains states: Wyoming—210, Nebraska— 
197, Montana—187, and North Dakota—148 (Stanford & Opler 1998). 

Data for county records were obtained from the following sources: 


(1) The collections at South Dakota State University, Brookings; Uni- 
versity of South Dakota, Vermillion; Northern State University, 
Aberdeen; and North Dakota State University, Fargo. 

(2) Private collections of Gerald Fauske, Fargo, N.D., who collected 
primarily in the vicinity of Sioux Falls; Bruce K. Harris, Clear 
Lake; Abran and Joseph Kean, Pierre; John S. Nordin, formerly 
of Webster, who collected extensively during 1965 to 1971; and 
the personal collection of the author. 

(3) A review of the literature, including Season Summaries of The 
Lepidopterists’ Society (1976-1993). 

(4) South Dakota collection records provided by non-resident col- 
lectors. 


Taxonomy and nomenclature follow A Catalogue/Checklist of the 
Butterflies of America North of Mexico (Miller & Brown 1981) and 
its supplement (Ferris 1989). 

The following species and subspecies have type localities designated 
in South Dakota: 


Erynnis persius fredericki—“‘Spearfish Canyon, Lawrence Co., South 
Dakota.” ; 


231 


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235 


VOLUME 48, NUMBER 3 


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JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


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VOLUME 48, NUMBER 3 Zon 


TABLE 2. Number of butterfly species documented for South Dakota counties. Coun- 
ties are listed alphabetically. 


Aurora 12 Hyde SP) 
Beadle sy Jackson 40 
Bennett 38 Jerauld Te 
Bon Homme 12 Jones 31 
Brookings 81 Kingsbury 22 
Brown 338 Lake 23 
Brule 46 Lawrence LN) 
Buffalo 155 Lincoln 35 
Butte 33 Lyman 37 
Campbell 15 McCook 23 
Charles Mix 38 McPherson 39 
Clark 16 Marshall 50 
Clay 42 Meade 74 
Codington 37 Mellette 38 
Corson 30 Miner 15 
Custer 106 Minnehaha 71 
Davison 30 Moody 23 
Day en Pennington ILIU7/ 
Deuel 52 Perkins 32 
Dewey al Potter 29 
Douglas 20 Roberts 64 
Edmunds 1 Sanborn 21 
Fall River 87 Shannon 43 
Faulk 20 Spink 18 
Grant 34 Stanley 51 
Gregory 30 Sully 33 
Haakon 24 Todd 46 
Hamlin 25 Tripp A7 
Hand 19 Turner 30 
Hanson Ny Union 49 
Harding 75 Walworth 26 
Hughes Sy) Yankton 49 
Hutchinson 23 Ziebach 26 


Hesperia dacotae—“‘Volga, South Dakota and Grinnell, Iowa.” 

Euchloe ausonides palaeoreios—‘‘Spearfish Canyon, Lawrence Co., 
South Dakota.” 

Colias alexandra krauthii—‘‘Black Hills, 12 mi. W. of Custer, Custer 
Co., South Dakota.” 

Everes amyntula valeriae—‘‘Lead, South Dakota.”’ 

Speyeria atlantis lurana—‘‘Harney Peak, Black Hills, South Dakota.” 


COUNTY RECORDS OF SOUTH DAKOTA BUTTERFLIES 


Superfamily Hesperioidea 
Family Hesperiidae 


1. Epargyreus clarus clarus (Cramer)—Brookings, Brule, Clay, Custer, Davison, Day, 
Deuel, Dewey, Fall River, Hand, Harding, Hughes, Lake, Lawrence, Meade, Mel- 
lette, Minnehaha, Pennington, Roberts, Shannon, Stanley, Union. 


ple 
12: 


13. 


14. 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


. Thorybes pylades (Scudder)—Custer, Fall River, Harding, Lawrence, Lincoln, Ly- 


man, Mellette, Pennington, Roberts, Yankton. 


. Staphylus hayhurstii (W. H. Edwards)—Yankton. 

. Erynnis icelus (Scudder and Burgess)—Custer, Lawrence, Pennington. 

. Erynnis brizo brizo (Boisduval and Le Conte)—Brookings, Minnehaha. 

. Erynnis juvenalis juvenalis (Fabricius)—Brookings, Charles Mix, Custer, Lawrence, 


Meade, Mellette, Pennington, Roberts, Todd. 


. Erynnis horatius (Scudder and Burgess)—Brookings, Day, Yankton. 
. Erynnis martialis (Scudder)—Lawrence. 
. Erynnis afranius (Lintner)—Butte, Custer, Fall River, Harding, Lawrence, Meade, 


Pennington, Shannon, Ziebach. 


. Erynnis persius fredericki H. A. Freeman—Custer, Harding, Lawrence, Meade, 


Pennington, Perkins. 

Pyrgus scriptura (Boisduval)—Custer, Fall River, Jackson. 

Pyrgus communis (Grote)—Beadle, Bennett, Brookings, Brown, Brule, Buffalo, Butte, 
Campbell, Charles Mix, Clay, Codington, Corson, Custer, Davison, Day, Fall River, 
Grant, Gregory, Haakon, Hanson, Harding, Hughes, Hutchinson, Hyde, Jackson, 
Jerauld, Jones, Lake, Lawrence, Lincoln, Lyman, McCook, McPherson, Meade, 
Mellette, Miner, Minnehaha, Pennington, Potter, Roberts, Sanborn, Shannon, Stan- 
ley, Sully, Todd, Tripp, Union, Walworth, Yankton. 

Pholisora catullus (Fabricius)—Bennett, Brookings, Brule, Butte, Charles Mix, Clay, 
Codington, Corson, Custer, Davison, Day, Dewey, Fall River, Faulk, Gregory, Haa- 
kon, Hanson, Harding, Hughes, Hutchinson, Jackson, Jones, Lawrence, McPherson, 
Meade, Mellette, Minnehaha, Pennington, Potter, Sanborn, Shannon, Stanley, Sully, 
Todd, Tripp, Turner, Yankton. 

Ancyloxypha numitor (Fabricius)—Bennett, Brookings, Clark, Clay, Codington, 
Day, Deuel, Fall River, Grant, Hamlin, Hughes, Jones, Kingsbury, Lake, Lawrence, 
Lincoln, Lyman, McCook, Minnehaha, Pennington, Roberts, Shannon, Stanley, Wal- 
worth, Yankton. 


. Oarisma powesheik (Parker)—Brookings, Brown, Codington, Day, Deuel, Grant, 


Hamlin, Marshall, Roberts. 


. Oarisma garita (Reakirt)—Butte, Custer, Fall River, Harding, Lawrence, Meade, 


Pennington. 


. Hylephila phyleus phyleus (Drury)—Clay. 
. Yvuretta rhesus (W. H. Edwards)—Bennett, Corson, Custer, Fall River, Harding, 


Jackson, Perkins. 


. Hesperia uncas uncas W. H. Edwards—Brookings, Codington, Custer, Day, Deuel, 


Fall River, Harding, Hughes, Jackson, Lawrence, McPherson, Meade, Pennington, 
Perkins, Potter, Stanley, Sully, Todd, Tripp. 


. Hesperia juba (Scudder)—Lawrence. 
. Hesperia comma assiniboia (Lyman)—Bennett, Custer, Day, Fall River, Harding, 


Lawrence, Marshall, McPherson, Pennington. 
Brookings, Corson, Day, Deuel, Fall River, Har- 
ding, Lawrence, Pennington, Tripp, Union, Ziebach. 


. Hesperia leonardus pawnee Dodge—Bennett, Brookings, Butte, Custer, Day, Fall 


River, Harding, Hughes, Jackson, Lawrence, Marshall, Meade, Mellette, Pennington, 
Perkins, Roberts, Shannon, Stanley, Tripp, Union. 


. Hesperia pahaska pahaska (Leussler)—Custer, Fall River, Harding, Lawrence, Pen- 


nington, Ziebach. 


. Hesperia dacotae (Skinner)—Brookings, Brown, Codington, Day, Deuel, Grant, 


Hamlin, Marshall, McPherson, Moody, Roberts. 


. Hesperia nevada (Scudder)—Pennington. 
. Polites peckius (W. Kirby)—Aurora, Brookings, Brown, Codington, Corson, Custer, 


Davison, Day, Deuel, Douglas, Hamlin, Hanson, Harding, Lawrence, Lincoln, Mar- 
shall, McPherson, Minnehaha, Moody, Pennington, Roberts, Tripp, Union, Yankton. 


. Polites themistocles (Latreille)—Aurora, Brookings, Brown, Brule, Clay, Codington, 


Custer, Davison, Day, Deuel, Fall River, Gregory, Hamlin, Hand, Hanson, Harding, 


VOLUME 48, NUMBER 3 | 239 


29. 


34. 


39. 


Hughes, Hyde, Jones, Lawrence, Lyman, Marshall, McPherson, Meade, Mellette, 
Minnehaha, Moody, Pennington, Roberts, Stanley, Tripp, Turner, Union, Yankton. 
Polites origenes rhena (W. H. Edwards)—Brookings, Codington, Custer, Day, Fall 
River, Harding, Lawrence, Marshall, Meade, Pennington, Shannon, Union. 

Polites mystic dacotah (W. H. Edwards)—Aurora, Bennett, Brookings, Brown, Clark, 
Codington, Custer, Day, Deuel, Douglas, Fall River, Hamlin, Kingsbury, Lake, 
Lawrence, Lincoln, Marshall, McPherson, Meade, Mellette, Minnehaha, Moody, 
Pennington, Roberts, Spink, Todd. 


. Wallengrenia egeremet (Scudder)—Brookings, Day, Deuel, Marshall, Roberts. 
. Pompeius verna (W. H. Edwards)—Hutchinson, Turner, Yankton. 


Atalopedes campestris campestris (Boisduval)—Bon Homme, Brookings, Brown, 
Charles Mix, Clay, Codington, Custer, Day, Deuel, Fall River, Grant, Gregory, 
Hughes, Jackson, Lake, Lawrence, Lincoln, Marshall, McPherson, Meade, Minne- 
haha, Roberts, Shannon, Spink, Stanley, Todd, Tripp, Union, Yankton. 

Atrytone arogos iowa (Scudder)—Brookings, Brown, Custer, Day, Deuel, Fall River, 
Gregory, Lawrence, McPherson, Meade, Roberts, Stanley, Todd, Tripp. 

Atrytone logan lagus (W. H. Edwards)—Brookings, Brule, Clay, Codington, Corson, 
Custer, Day, Deuel, Douglas, Edmunds, Fall River, Grant, Gregory, Hamlin, Hand, 
Harding, Hughes, Hutchinson, Jerauld, Kingsbury, Marshall, McCook, McPherson, 
Meade, Mellette, Minnehaha, Moody, Pennington, Perkins, Potter, Roberts, Spink, 
Tripp, Union, Walworth, Yankton, Ziebach. 


. Ochlodes sylvanoides napa (W. H. Edwards)—Butte, Lawrence, Meade. 
. Poanes massasoit massasoit (Scudder)—Brookings, Roberts. 
. Poanes hobomok hobomok (Harris)—Brookings, Brule, Charles Mix, Custer, Davi- 


son, Day, Deuel, Douglas, Fall River, Haakon, Harding, Hughes, Jerauld, Lake, 
Lawrence, Lyman, Marshall, McCook, Mellette, Minnehaha, Pennington, Roberts, 
Stanley, Tripp, Turner, Union. 


. Poanes zabulon (Boisduval and Le Conte)—Brule. 

. Poanes taxiles (W. H. Edwards)—Custer, Fall River, Lawrence, Meade, Pennington. 
. Poanes viator viator (W. H. Edwards)—Brookings, Day, Deuel, Roberts. 

. Euphyes vestris kiowah (Reakirt)—Brookings, Brule, Custer, Day, Fall River, Har- 


ding, Lawrence, Marshall, Minnehaha, Pennington, Perkins, Roberts, Stanley, Sully, 
Todd, Tripp, Walworth. 


43a.Atrytonopsis hianna ssp.—Day, Marshall, McPherson, Roberts. 
43b.Atrytonopsis hianna turneri H. A. Freeman—Bennett, Fall River, Haakon, Harding, 


44. 
45. 


46. 


47. 
48. 


Jones, Lyman, Meade, Mellette, Pennington, Perkins, Shannon. 

Amblyscirtes simius W. H. Edwards—Custer, Fall River, Meade, Pennington. 
Amblyscirtes oslari (Skinner)—Custer, Fall River, Haakon, Harding, Lawrence, 
Meade, Mellette, Pennington. 

Amblyscirtes vialis (W. H. Edwards)—Brookings, Charles Mix, Custer, Day, Fall 
River, Gregory, Haakon, Harding, Hughes, Hutchinson, Jackson, Jones, Lawrence, 
Lincoln, Meade, Mellette, Pennington, Perkins, Sanborn, Shannon, Sully, Todd, 
Tripp, Walworth. 

Lerodea eufala (W. H. Edwards)—Day, Minnehaha. 

Megathymus streckeri leussleri Holland—Custer, Fall River, Harding, Jackson, 
Lawrence, Meade, Pennington, Stanley, Todd, Ziebach. 


Superfamily Papilionoidea 


Family Papilionidae 


. Parnassius phoebus sayii W. H. Edwards—Custer, Lawrence, Meade, Pennington. 
. Battus philenor philenor (Linnaeus)—Brookings, Minnehaha. 

. Eurytides marcellus (Cramer)—Minnehaha. 

. Papilio polyxenes asterius Stoll—Bennett, Bon Homme, Brookings, Brown, Brule, 


Charles Mix, Custer, Davison, Day, Deuel, Fall River, Grant, Haakon, Hand, Hughes, 
Lawrence, Marshall, Minnehaha, Potter, Roberts, Stanley, Turner, Union, Yankton. 


240 


53. 
OA. 


D0. 
36. 


o7. 


58. 


59. 
60. 


61. 
62. 


63. 


64. 


65. 


66. 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Papilio bairdii bairdii W. H. Edwards—Custer, Fall River, Harding, Pennington, 
Shannon, Stanley. 

Papilio zelicaon nitra W. H. Edwards—Butte, Custer, Fall River, Harding, Law- 
rence, McPherson, Meade, Pennington, Perkins, Shannon, Ziebach. 

Papilio indra indra Reakirt—Custer, Fall River, Lawrence, Pennington. 
Heraclides cresphontes (Cramer)—Brookings, Codington, Lincoln, Minnehaha, 
Union, Yankton. 

Pterourus glaucus glaucus (Linnaeus)—Brookings, Brown, Brule, Charles Mix, Cod- 
ington, Davison, Day, Deuel, Douglas, Faulk, Gregory, Hamlin, Hand, Hughes, 
Hutchinson, Hyde, Jerauld, Kingsbury, Lake, Lincoln, Lyman, Marshall, McCook, 
Mellette, Miner, Minnehaha, Moody, Roberts, Spink, Stanley, Tripp, Turner, Union, 
Walworth, Yankton. 

Pterourus canadensis (Rothschild and Jordan)—Custer, Fall River, Lawrence, Meade, 
Pennington, Roberts. 

Pterourus rutulus rutulus (Lucas)—Custer, Fall River, Lawrence, Pennington. 
Pterourus multicaudatus (W. F. Kirby)—Brule, Custer, Fall River, Harding, Law- 
rence, Meade, Pennington, Shannon, Todd. 

Pterourus eurymedon (Lucas)—Lawrence, Pennington. 

Pterourus troilus troilus (Linnaeus)—Minnehaha, Pennington. 


Family Pieridae 


Neophasia menapia menapia (C. and R. Felder)—Custer, Fall River, Lawrence, 
Meade, Pennington. 

Pontia sisymbrii nordini (K. Johnson)—Custer, Fall River, Harding, Lawrence, 
Pennington. 

Pontia protodice (Boisduval and Le Conte)—Beadle, Bennett, Brookings, Brule, 
Buffalo, Butte, Charles Mix, Clay, Corson, Custer, Davison, Day, Douglas, Fall River, 
Gregory, Hand, Harding, Hughes, Hutchinson, Hyde, Jackson, Jones, Lawrence, 
Lincoln, Lyman, McCook, McPherson, Meade, Miner, Minnehaha, Pennington, Pot- 
ter, Sanborn, Shannon, Spink, Stanley, Sully, Tripp, Turner, Union, Yankton. 
Pontia occidentalis occidentalis (Reakirt)—Bennett, Brule, Butte, Corson, Custer, 
Day, Fall River, Hughes, Hyde, Lawrence, Lyman, McPherson, Mellette, Penning- 
ton, Sanborn, Stanley, Sully, Walworth, Ziebach. 


67a. Pieris napi oleracea Harris—Marshall, Roberts. 


67b.Pieris napi mcdunnoughi Remington 


68. 


C2, 
73. 


Lawrence. 

Pieris rapae (Linnaeus)—Aurora, Bennett, Bon Homme, Brookings, Brown, Brule, 
Buffalo, Campbell, Charles Mix, Clark, Clay, Codington, Corson, Custer, Davison, 
Day, Deuel, Dewey, Douglas, Edmunds, Fall River, Faulk, Grant, Gregory, Haakon, 
Hamlin, Hand, Hanson, Harding, Hughes, Hutchinson, Hyde, Jackson, Jerauld, 
Jones, Kingsbury, Lake, Lawrence, Lincoln, Lyman, Marshall, McCook, McPherson, 
Meade, Mellette, Miner, Minnehaha, Moody, Pennington, Potter, Roberts, Sanborn, 
Shannon, Spink, Stanley, Sully, Todd, Tripp, Turner, Union, Walworth, Yankton. 


. Ascia monuste ssp.—Brown. 
. Euchloe ausonides palaeoreios K. Johnson—Custer, Lawrence, Meade, Pennington. 
. Euchloe olympia W. H. Edwards—Bennett, Brookings, Brule, Butte, Campbell, 


Clay, Corson, Custer, Dewey, Fall River, Haakon, Harding, Hughes, Jackson, Jones, 
Lawrence, Lyman, Marshall, McPherson, Meade, Mellette, Minnehaha, Pennington, 
Perkins, Potter, Shannon, Stanley, Sully, Todd, Union, Yankton, Ziebach. 
Anthocharis sara julia W. H. Edwards—Lawrence. 

Colias philodice ssp.—Beadle, Bennett, Brookings, Brown, Brule, Buffalo, Butte, 
Campbell, Charles Mix, Clark, Clay, Codington, Corson, Custer, Day, Deuel, Dewey, 
Douglas, Edmunds, Fall River, Faulk, Grant, Haakon, Hamlin, Hand, Hanson, 
Harding, Hughes, Hutchinson, Hyde, Jackson, Jerauld, Jones, Kingsbury, Lake, 
Lawrence, Lincoln, Lyman, Marshall, McCook, McPherson, Meade, Mellette, Miner, 
Minnehaha, Moody, Pennington, Perkins, Potter, Roberts, Sanborn, Shannon, Spink, 
Stanley, Sully, Todd, Tripp, Turner, Union, Walworth, Yankton, Ziebach. 


VOLUME 48, NUMBER 3 : . 241 


7A. 


Colias eurytheme Boisduval—Aurora, Beadle, Bennett, Bon Homme, Brookings, 
Brown, Brule, Buffalo, Butte, Campbell, Charles Mix, Clark, Clay, Codington, Cor- 
son, Custer, Davison, Day, Deuel, Dewey, Douglas, Edmunds, Fall River, Faulk, 
Grant, Gregory, Haakon, Hamlin, Hand, Hanson, Harding, Hughes, Hutchinson, 
Hyde, Jackson, Jerauld, Jones, Kingsbury, Lake, Lawrence, Lincoln, Lyman, Mar- 
shall, McCook, McPherson, Meade, Mellette, Miner, Minnehaha, Moody, Penning- 
ton, Perkins, Potter, Roberts, Sanborn, Shannon, Spink, Stanley, Sully, Todd, Tripp, 
Turner, Union, Walworth, Yankton, Ziebach. 


75a.Colias alexandra alexandra W. H. Edwards—Custer, Fall River, Jackson, Penning- 


ton. 


75b.Colias alexandra krauthii Klots—Custer, Fall River, Lawrence, Pennington. 


io: 


CH 


88. 


89. 


90. 


oe 


o2: 


93. 


94. 
95. 


Zerene cesonia cesonia (Stoll)—Brookings, Butte, Charles Mix, Corson, Fall River, 
Gregory, Hughes, Minnehaha, Pennington, Roberts, Sanborn, Stanley, Union, Yank- 
ton. 


. Phoebis sennae eubule (Linnaeus)—Clay, Minnehaha, Union. 

. Phoebis agarithe agarithe (Boisduval)—Brookings, Minnehaha. 

. Eurema daira daira (Godart)—Minnehaha. 

. Eurema mexicanum (Boisduval)—Corson, Custer, Minnehaha, Pennington. 

. Eurema lisa lisa (Boisduval and Le Conte)—Brookings, Clay, Day, Hughes, Min- 


nehaha, Union, Yankton. 


. Eurema nicippe (Cramer)—Minnehaha. 


Nathalis iole Boisduval—Bon Homme, Brookings, Charles Mix, Clay, Custer, Day, 
Deuel, Fall River, Gregory, Hughes, Jackson, Lawrence, Lyman, Minnehaha, Rob- 
erts, Sanborn, Stanley, Turner, Union, Yankton. 


Family Lycaeidae 


. Feniseca tarquinius tarquinius (Fabricius)—Roberts. 
. Lycaena phlaeas americana Harris—Brown, Jackson, Meade. 
. Gaeides xanthoides dione (Scudder)—Bennett, Brookings, Brule, Buffalo, Charles 


Mix, Clay, Corson, Custer, Day, Deuel, Douglas, Fall River, Grant, Gregory, Har- 
ding, Hughes, Jackson, Jones, Lawrence, Lyman, Marshall, McPherson, Meade, 
Minnehaha, Pennington, Perkins, Potter, Shannon, Stanley, Sully, Todd, Tripp. 
Hyllolycaena hyllus (Cramer)—Beadle, Bennett, Brookings, Brule, Charles Mix, 
Clark, Clay, Codington, Corson, Custer, Davison, Day, Deuel, Edmunds, Fall River, 
Faulk, Grant, Gregory, Hamlin, Hand, Hanson, Hughes, Hutchinson, Hyde, Jackson, 
Jerauld, Jones, Kingsbury, Lawrence, Lyman, Marshall, McPherson, Meade, Mel- 
lette, Miner, Minnehaha, Moody, Pennington, Potter, Roberts, Shannon, Spink, Sully, 
Todd, Tripp, Turner, Union, Yankton. 

Chalceria rubida longi (K. Johnson and Balogh)—Butte, Custer, Harding, Lawrence, 
Meade, Pennington. 

Epidemia helloides (Boisduval)—Bennett, Brookings, Brule, Butte, Clark, Custer, 
Day, Dewey, Haakon, Harding, Hughes, Jackson, Jones, Lawrence, Marshall, Meade, 
Mellette, Minnehaha, Pennington, Potter, Roberts, Shannon, Stanley, Todd, Tripp. 
Harkenclenus titus titus (Fabricius)—Bennett, Bon Homme, Brookings, Brule, Charles 
Mix, Codington, Day, Deuel, Fall River, Grant, Gregory, Harding, Hughes, Law- 
rence, Marshall, Meade, Pennington, Potter, Roberts, Stanley, Sully, Todd, Tripp, 
Union, Walworth, Ziebach. 

Satyrium acadicum montanensis (Watson and W. P. Comstock)—Beadle, Brook- 
ings, Brown, Kingsbury, Lawrence, Lyman, McCook, Meade, Pennington, Todd, 
Tripp. 

Satyrium edwardsii (Grote and Robinson)—Roberts. 

Satyrium calanus falacer (Godart)—Brule, Charles Mix, Day, Deuel, Lawrence, 
Meade, Minnehaha, Roberts, Sully, Tripp, Union. 

Satyrium caryaevorum (McDunnough)—Lawrence. 

Satyrium liparops ssp.—Day, Harding, Hughes, Jackson, Lawrence, Lyman, Min- 
nehaha, Pennington. 


242 


104. 


105. 


106. 


107. 


108. 


109. 


EO! 
JEU. 
U2. 
113. 


114. 


JUILS). 


116. 


JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Mitoura siva siva (W. H. Edwards)—Brule, Custer, Fall River, Haakon, Harding, 
Hughes, Jackson, Jones, Lyman, Meade, Mellette, Pennington, Shannon, Ziebach. 


. Mitoura grynea grynea (Hubner)—Union. 

. Incisalia augustinus iroides (Boisduval)—Custer, Lawrence, Meade, Pennington. 

. Incisalia polia obscura Ferris and Fisher—Custer, Lawrence, Pennington. 

. Incisalia eryphon eryphon (Boisduval)—Custer, Fall River, Harding, Lawrence, 


Meade, Pennington, Todd. 


. Strymon melinus atrofasciatus McDunnough—Bennett, Bon Homme, Brookings, 


Brule, Butte, Charles Mix, Clay, Custer, Davison, Day, Fall River, Gregory, Haakon, 
Harding, Hughes, Jackson, Lawrence, Lyman, McCook, Meade, Mellette, Minne- 
haha, Pennington, Shannon, Stanley, Sully, Todd, Tripp, Union, Yankton, Ziebach. 
Leptotes marina (Reakirt)—Minnehaha, Pennington. 

Hemiargus isola alce (W. H. Edwards)—Brookings, Butte, Clay, Codington, Custer, 
Day, Fall River, Haakon, Hamlin, Hanson, Harding, Hughes, Jones, Lawrence, 
Lyman, Marshall, Meade, Minnehaha, Pennington, Stanley, Sully, Todd, Tripp, 
Union, Yankton, Ziebach. 

Everes comyntas comyntas (Godart)—Brookings, Brown, Campbell, Charles Mix, 
Clay, Codington, Custer, Davison, Day, Faulk, Grant, Gregory, Haakon, Hamlin, 
Harding, Hughes, Jackson, Lake, Lawrence, Lincoln, Lyman, Marshall, McCook, 
McPherson, Mellette, Minnehaha, Moody, Pennington, Potter, Roberts, Sully, Todd, 
Tripp, Union, Walworth, Yankton. 

Everes amyntula valeriae Clench—Custer, Fall River, Harding, Lawrence, Pen- 
nington. 

Celastrina argiolus ssp.—Beadle, Brookings, Brule, Clay, Codington, Custer, Dav- 
ison, Day, Deuel, Dewey, Douglas, Fall River, Hanson, Harding, Hughes, Jerauld, 
Kingsbury, Lake, Lawrence, Lincoln, Marshall, McPherson, Meade, Minnehaha, 
Moody, Pennington, Roberts, Shannon, Stanley, Union, Yankton. 

Glaucopsyche piasus daunia (W.H. Edwards)—Custer, Harding, Lawrence, Pen- 
nington. 

Glaucopsyche lygdamus oro (Scudder)—Butte, Campbell, Corson, Custer, Day, 
Dewey, Fall River, Harding, Hughes, Jackson, Jones, Lawrence, Marshall, Meade, 
Mellette, Pennington, Perkins, Shannon, Stanley, Sully. 

Lycaeides melissa melissa (W. H. Edwards)—Aurora, Beadle, Bennett, Brookings, 
Brown, Brule, Buffalo, Butte, Campbell, Charles Mix, Clark, Clay, Codington, Cor- 
son, Custer, Davison, Day, Deuel, Dewey, Douglas, Edmunds, Fall River, Faulk, 
Grant, Gregory, Haakon, Hamlin, Hanson, Harding, Hughes, Hyde, Jackson, Jerauld, 
Jones, Kingsbury, Lake, Lawrence, Lincoln, Lyman, Marshall, McCook, McPherson, 
Meade, Mellette, Miner, Minnehaha, Moody, Pennington, Perkins, Potter, Roberts, 
Sanborn, Shannon, Spink, Stanley, Sully, Todd, Tripp, Union, Walworth, Yankton, 
Ziebach. 

Plebejus saepiolus saepiolus (Boisduval)—Custer, Fall River, Harding, Lawrence, 
Meade, Pennington. 

Icaricia icarioides lycea (W. H. Edwards)—Custer, Harding, Lawrence, Pennington. 
Icaricia shasta minnehaha (Scudder)-—Lawrence, Pennington. 

Icaricia acmon lutzi (dos Passos)—Bennett, Butte, Custer, Day, Fall River, Harding, 
Jackson, Lawrence, Meade, Mellette, Pennington, Perkins, Todd. 

Agriades rusticus rusticus (W. H. Edwards)—Custer, Lawrence, Pennington. 


Family Riodinidae 


Apodemia mormo mormo (C. and R. Felder)—Pennington. 


Family Libytheidae 


Libytheana bachmanii bachmanii (Kirkland)—Brookings, Clay, Davison, Day, Min- 
nehaha, Yankton. 


VOLUME 48, NUMBER 3 | 243 


Family Heliconiidae 


117. Agraulis vanillae incarnata (Riley)—Butte, Minnehaha, Pennington. 


Family Nymphalidae 


118. Euptoieta claudia (Cramer)—Bennett, Bon Homme, Brookings, Brown, Brule, Buf- 
falo, Butte, Campbell, Charles Mix, Clay, Codington, Custer, Davison, Day, Deuel, 
Dewey, Douglas, Fall River, Faulk, Grant, Hamlin, Hand, Harding, Hughes, Jackson, 
Jones, Kingsbury, Lawrence, Lincoln, Lyman, Marshall, McCook, McPherson, Meade, 
Mellette, Minnehaha, Moody, Pennington, Perkins, Potter, Roberts, Sanborn, Shan- 
non, Spink, Stanley, Sully, Todd, Tripp, Turner, Union, Walworth, Yankton, Ziebach. 

119. Speyeria cybele cybele (Fabricius)—Brookings, Brule, Clay, Corson, Custer, Day, 
Deuel, Dewey, Faulk, Grant, Gregory, Hamlin, Harding, Hutchinson, Hyde, Jack- 
son, Jones, Lawrence, Lincoln, Lyman, Marshall, McCook, Meade, Miner, Minne- 
haha, Pennington, Perkins, Potter, Roberts, Stanley, Sully, Todd, Tripp, Turner, 
Union, Walworth, Yankton. 

120. Speyeria aphrodite alcestris (W. H. Edwards)—Brookings, Codington, Corson, Cus- 
ter, Day, Deuel, Fall River, Grant, Haakon, Harding, Hughes, Lawrence, Marshall, 
McPherson, Meade, Minnehaha, Pennington, Perkins, Roberts, Shannon, Union, 
Walworth, Ziebach. 

121. Speyeria idalia (Drury)—Beadle, Bennett, Bon Homme, Brookings, Brown, Brule, 
Campbell, Charles Mix, Clay, Codington, Corson, Custer, Davison, Day, Deuel, 
Douglas, Fall River, Faulk, Grant, Gregory, Haakon, Hamlin, Hanson, Hughes, 
Hutchinson, Hyde, Jerauld, Jones, Kingsbury, Lake, Lawrence, Lincoln, Lyman, 
Marshall, McCook, McPherson, Meade, Minnehaha, Moody, Pennington, Potter, 
Roberts, Sanborn, Shannon, Spink, Stanley, Sully, Todd, Tripp, Turner, Union, 
Walworth, Yankton, Ziebach. 

122. Speyeria edwardsii (Reakirt)—Bennett, Brule, Butte, Custer, Dewey, Fall River, 
Harding, Hughes, Jackson, Jones, Lawrence, Lyman, McPherson, Meade, Mellette, 
Pennington, Shannon, Stanley, Sully, Todd. 

123. Speyeria coronis ssp.—Custer, Fall River, Harding, Lawrence, Pennington. 

124. Speyeria zerene garretti (Gunder)—Custer, Harding, Lawrence, Pennington. 

125. Speyeria callippe calgariana (McDunnough)—Campbell, Custer, Fall River, Har- 
ding, Lawrence, McPherson, Meade, Pennington, Perkins, Ziebach. 

126a.Speyeria atlantis ssp.—Custer, Lawrence, Pennington. 

126b.Speyeria atlantis lurana dos Passos and Grey—Custer, Lawrence, Meade, Penning- 
ton. 

127. Speyeria mormonia eurynome (W. H. Edwards)—Custer, Lawrence, Pennington, 
Shannon. 

128a.Clossiana selene myrina (Cramer)—Brookings, Brown, Codington, Day, Deuel, 
Grant, Marshall, McPherson, Roberts. 

128b.Clossiana selene nebraskensis (Holland)—Bennett, Mellette, Todd. 

128c.Clossiana selene sabulocollis (Kohler)—Custer, Lawrence, Pennington. 

129. Clossiana bellona toddi (Holland)—Brookings, Brown, Clay, Day, Deuel, Grant, 
Kingsbury, Marshall, Miner, Minnehaha, Moody, Roberts. 

130. Charidryas gorgone carlota (Reakirt)—Beadle, Bennett, Brookings, Brown, Brule, 
Buffalo, Butte, Charles Mix, Corson, Custer, Day, Dewey, Fall River, Faulk, Haakon, 
Harding, Hughes, Hutchinson, Hyde, Jones, Lawrence, Lyman, Marshall, McPher- 
son, Meade, Mellette, Minnehaha, Pennington, Perkins, Shannon, Spink, Stanley, 
Todd, Tripp, Union, Yankton, Ziebach. 

131. Charidryas nycteis nycteis (Doubleday and Hewitson)—Bennett, Brookings, Brule, 
Clay, Custer, Day, Deuel, Dewey, Gregory, Hyde, Lawrence, Marshall, Minnehaha, 
Pennington, Roberts, Todd, Yankton. 

132. Charidryas acastus (W. H. Edwards)—Brule, Custer, Harding, Hughes, Stanley. 

133. Anthanassa texana texana (W. H. Edwards)—Roberts. 

134. Phyciodes tharos tharos (Drury)—Beadle, Bennett, Brookings, Brown, Brule, Buf- 


244 


135. 
136. 
137. 
138. 
139. 


140. 


141. 


142. 
143. 
144. 
145. 
146. 


147. 
148. 


149. 


150. 


151. 


152. 
153. 


154. 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


falo, Butte, Campbell, Charles Mix, Clay, Codington, Corson, Custer, Davison, Day, 
Deuel, Dewey, Douglas, Fall River, Faulk, Grant, Gregory, Haakon, Harding, Hughes, 
Hutchinson, Hyde, Jackson, Jones, Kingsbury, Lake, Lawrence, Lincoln, Lyman, 
Marshall, McCook, McPherson, Meade, Mellette, Miner, Minnehaha, Pennington, 
Perkins, Potter, Roberts, Sanborn, Shannon, Spink, Stanley, Sully, Todd, Tripp, 
Turner, Union, Walworth, Yankton, Ziebach. 

Phyciodes pascoensis W. G. Wright—Brookings, Custer, Day, Deuel, Grant, Har- 
ding, Jackson, Lawrence, Marshall, Pennington, Roberts. 

Phyciodes batesii (Reakirt)—Custer, Lawrence, Meade, Pennington. 

Phyciodes pratensis camillus W. H. Edwards—Fall River, Lawrence. 

Phyciodes pallidus barnesi Skinner—Custer, Fall River, Lawrence. 

Euphydryas anicia bernadetta Leussler—Butte, Custer, Fall River, Harding, Law- 
rence, Pennington. 

Polygonia interrogationis (Fabricius)—Bon Homme, Brookings, Brule, Charles Mix, 
Clay, Davison, Day, Deuel, Faulk, Grant, Hughes, Hyde, Lake, Lincoln, Marshall, 
Meade, Mellette, Minnehaha, Pennington, Roberts, Stanley, Todd, Turner, Union, 
Walworth, Yankton. 

Polygonia comma (Harris)—Brookings, Charles Mix, Clay, Custer, Day, Deuel, 
Dewey, Gregory, Hand, Hughes, Hutchinson, Hyde, Lincoln, Lyman, Marshall, 
Minnehaha, Potter, Roberts, Turner, Walworth, Yankton. 

Polygonia satyrus (W. H. Edwards)—Custer, Lawrence, Pennington. 

Polygonia faunus ssp.—Custer, Fall River, Lawrence, Meade, Pennington. 
Polygonia zephryus (W. H. Edwards)—Custer, Fall River, Lawrence, Meade, Pen- 
nington. 

Polygonia progne progne (Cramer)—Brookings, Brule, Custer, Day, Fall River, 
Harding, Lawrence, Lincoln, Minnehaha, Pennington, Roberts, Yankton. 
Nymphalis vau-album j-album (Boisduval and Le Conte)—Brookings, Custer, Law- 
rence, Marshall, Meade, Pennington, Roberts. 

Nymphalis californica californica (Boisduval)—Brookings, Lawrence. 

Nymphalis antiopa antiopa (Linnaeus)—Bennett, Brookings, Brown, Brule, Butte, 
Charles Mix, Clark, Clay, Codington, Corson, Custer, Davison, Day, Deuel, Fall 
River, Faulk, Grant, Hand, Harding, Hughes, Hutchinson, Jackson, Jerauld, Jones, 
Lake, Lawrence, Lincoln, Lyman, Marshall, McPherson, Meade, Mellette, Miner, 
Minnehaha, Moody, Pennington, Perkins, Potter, Roberts, Shannon, Stanley, Sully, 
Todd, Tripp, Turner, Union, Yankton, Ziebach. 

Aglais milberti milberti (Godart)—Brookings, Custer, Day, Deuel, Fall River, Har- 
ding, Hughes, Lawrence, Meade, Minnehaha, Pennington, Perkins, Shannon, Stan- 
ley. 

Vanessa virginiensis (Drury)—Brookings, Butte, Codington, Custer, Day, Deuel, 
Edmunds, Fall River, Jackson, Lawrence, Meade, Minnehaha, Pennington, Stanley, 
Turner, Union, Yankton. 

Vanessa cardui (Linnaeus)—Aurora, Beadle, Bennett, Brookings, Brown, Brule, 
Buffalo, Butte, Campbell, Charles Mix, Clark, Clay, Codington, Corson, Custer, 
Davison, Day, Deuel, Dewey, Edmunds, Fall River, Faulk, Grant, Gregory, Haakon, 
Hamlin, Hand, Hanson, Harding, Hughes, Hyde, Jackson, Jerauld, Jones, Kingsbury, 
Lake, Lawrence, Lincoln, Lyman, Marshall, McCook, McPherson, Meade, Mellette, 
Miner, Minnehaha, Pennington, Perkins, Potter, Roberts, Sanborn, Shannon, Spink, 
Stanley, Sully, Todd, Tripp, Turner, Union, Walworth, Yankton, Ziebach. 
Vanessa annabella (Field)—Pennington. 

Vanessa atalanta rubria (Fruhstorfer)—Beadle, Bennett, Brookings, Brown, Brule, 
Butte, Charles Mix, Clark, Clay, Codington, Corson, Custer, Davison, Day, Deuel, 
Dewey, Douglas, Edmunds, Fall River, Faulk, Grant, Gregory, Haakon, Hamlin, 
Hand, Hanson, Harding, Hughes, Hutchinson, Hyde, Jackson, Jones, Kingsbury, 
Lake, Lawrence, Lincoln, Lyman, Marshall, McPherson, Meade, Mellette, Minne- 
haha, Moody, Pennington, Perkins, Potter, Roberts, Sanborn, Shannon, Spink, Stan- 
ley, Sully, Todd, Tripp, Turner, Union, Walworth, Yankton, Ziebach. 

Junonia coenia Hiibner—Aurora, Brookings, Brown, Brule, Charles Mix, Clay, Cod- 


VOLUME 48, NUMBER 3 245 


155a. 
155b. 


155c. 


156. 


hoe 
158. 


159. 
160. 


161. 


162. 


163. 


164. 


165a. 


165b. 


166. 


167. 
168. 
169. 


170. 
vel. 


ington, Custer, Day, Deuel, Edmunds, Lincoln, Minnehaha, Tripp, Turner, Union, 
Yankton. 

Basilarchia arthemis arthemis (Drury)—Brookings, Day, Marshall, Roberts. 
Basilarchia arthemis rubrofasciata Barnes and McDunnough—Brookings, Brown, 
Day, Deuel, Marshall, Minnehaha, Roberts. 

Basilarchia arthemis astyanax (Fabricius)—Brookings, Charles Mix, Clark, Clay, 
Codington, Davison, Day, Deuel, Grant, Hamlin, Hand, Hughes, Kingsbury, Lin- 
coln, Marshall, Minnehaha, Moody, Roberts, Stanley, Turner, Union, Yankton. 
Basilarchia archippus archippus (Cramer)—Beadle, Bennett, Brookings, Brown, 
Brule, Buffalo, Charles Mix, Clark, Clay, Davison, Day, Deuel, Douglas, Fall River, 
Grant, Hand, Hanson, Hughes, Hutchinson, Hyde, Jackson, Jones, Lake, Lawrence, 
Lincoln, Lyman, Marshall, McCook, Miner, Minnehaha, Moody, Pennington, Potter, 
Roberts, Sanborn, Shannon, Stanley, Sully, Todd, Turner, Union, Yankton. 
Basilarchia weidemeyerii oberfoelli (F. M. Brown)—Custer, Fall River, Harding, 
Jackson, Lawrence, Meade, Pennington, Perkins, Shannon, Todd. 

Mestra amymone (Ménetriés)—Brookings. 

Anaea andria Scudder—Shannon, Todd, Tripp. 

Asterocampa celtis celtis (Boisduval and Le Conte)—Brookings, Brule, Buffalo, 
Charles Mix, Clay, Davison, Day, Fall River, Grant, Hughes, Hutchinson, Kingsbury, 
Lincoln, Marshall, McCook, Minnehaha, Roberts, Stanley, Todd, Tripp, Turner, 
Union, Walworth, Yankton. 

Asterocampa clyton clyton (Boisduval and Le Conte)—Brookings, Day, Kingsbury, 
Lincoln, Minnehaha, Yankton. 


Family Satyridae 


Enodia anthedon A. H. Clark—Day, Deuel, Lincoln, Marshall, Minnehaha, Roberts, 
Tripp. 

Satyrodes eurydice ssp.—Bennett, Brookings, Corson, Day, Deuel, Grant, Hamlin, 
Hughes, Marshall, McCook, Moody, Roberts, Todd, Tripp. 

Megisto cymela cymela (Cramer)—Aurora, Bennett, Bon Homme, Brookings, Brule, 
Charles Mix, Clark, Clay, Corson, Day, Deuel, Douglas, Faulk, Gregory, Harding, 
Hughes, Hutchinson, Jerauld, Lawrence, Lincoln, Marshall, Minnehaha, Roberts, 
Sanborn, Stanley, Sully, Tripp, Turner, Union, Yankton. 

Coenonympha tullia ssp.—Aurora, Bennett, Brookings, Brule, Charles Mix, Cod- 
ington, Corson, Davison, Day, Deuel, Dewey, Grant, Gregory, Haakon, Hamlin, 
Hughes, Jackson, Jerauld, Jones, Lake, Lyman, Marshall, McPherson, Mellette, Min- 
nehaha, Perkins, Potter, Roberts, Sanborn, Shannon, Stanley, Sully, Todd, Tripp, 
Turner, Walworth, Ziebach. 

Coenonympha tullia benjamini McDunnough—Butte, Custer, Fall River, Harding, 
Lawrence, Meade, Pennington. 

Cercyonis pegala spp.—Aurora, Bennett, Bon Homme, Brookings, Brown, Brule, 
Buffalo, Butte, Campbell, Charles Mix, Clark, Clay, Codington, Corson, Custer, 
Davison, Day, Deuel, Dewey, Douglas, Edmunds, Fall River, Faulk, Grant, Gregory, 
Hamlin, Hand, Hanson, Harding, Hughes, Hutchinson, Hyde, Jackson, Jerauld, 
Jones, Kingsbury, Lake, Lawrence, Lincoln, Lyman, Marshall, McCook, McPherson, 
Meade, Mellette, Minnehaha, Moody, Pennington, Perkins, Potter, Roberts, Sanborn, 
Shannon, Spink, Stanley, Sully, Todd, Tripp, Turner, Union, Walworth, Yankton, 
Ziebach. 

Cercyonis meadii meadii (W. H. Edwards)—Fall River, Harding, Lawrence, Meade, 
Pennington. 

Cercyonis oetus charon (W. H. Edwards)—Custer, Harding, Lawrence, Meade, 
Pennington, Perkins. 

Neominois ridingsii ridingsii (W. H. Edwards)—Pennington. 

Oeneis chryxus chryxus (Doubleday and Hewitson)—Pennington. 

Oeneis uhleri ssp.—Brookings, Brown, Butte, Custer, Day, Fall River, Harding, 
Lawrence, Marshall, McPherson, Pennington, Perkins, Potter, Roberts, Shannon. 


246 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Family Danaidae 


172. Danaus plexippus (Linnaeus)—Aurora, Beadle, Bennett, Brookings, Brown, Brule, 
Buffalo, Campbell, Charles Mix, Clark, Clay, Codington, Custer, Davison, Day, 
Deuel, Douglas, Fall River, Faulk, Grant, Gregory, Hamlin, Hand, Hanson, Harding, 
Hughes, Hutchinson, Hyde, Jecicon Jerauld, Jones, Kingsbary, Lake, Lawrence, 
Lincoln, Lyman, Marshall, McCook, McPherson, Miner, Minnehaha, Moody, Pen- 
nington, Perkins, Potter, Roberts, Sanborn, Spink, Stanley, Sully, Todd, Tripp, Tur- 
ner,Union, Walworth, Yankton. 


ACKNOWLEDGMENTS 


I acknowledge with sincere gratitude the assistance of Clifford D. Ferris, Paul A. Opler, 
and Ray E. Stanford for determination of specimens, distribution records, and reviewing 
portions of the manuscript. Special thanks to my young collecting companions, Abran 
and Joseph Kean, for their dedicated effort in obtaining numerous county and state records. 
Access to university collections was made possible by Burruss McDaniel, South Dakota 
State University, and James Schmulbach, University of South Dakota. Many additional 
records were kindly provided by the following: C. V. Covell, D. L. Eiler, G. Fauske, B. 
K. Harris, S. J. Kohler, W. Lacey, W. W. McGuire, J. S. Nordin, J. Oberfoell, R. L. 
Patterson, B. Preheim, J. M. Reiser, R. C. Rosche, K. Roever, R. A. Royer, D. Skadsen, 
D. Schmidt, S. M. Spomer, and E. G. Unkenholz. 


LITERATURE CITED 


Conway, P. J. 1966. A quick collecting trip to the Black Hills. Newsl. Assoc. Minn. 
Entomol. 1:16-19. 

FERRIS, C. D. 1969. Two new forms of Plebejinae from Wyoming. J. Res. Lepid. 8:91- 
93. 


1976. A proposed revision of non-arctic Parnassius phoebus Fabricius in North 

America (Papilionidae). J. Res. Lepid. 15:1-22. 

1981. Field study of Phyciodes batesii (Reakirt) and P. tharos (Drury) from a 

site in the Black Hills, South Dakota (Lepidoptera: Nymphalidae: Melitaeinae). J. 

Res. Lepid. 20:285-289. 

1988. Speyeria atlantis phenotypes in the Southern Rocky Mountains (Lepi- 

doptera: Nymphalidae: Argynninae). J. Res. Lepid. 22:101-114. 

. ed. 1989. Supplement to: a catalogue/checklist of the butterflies of America 
North of Mexico. Mem. Lep. Soc. 1:1-145. 

FERRIS, C. D. & F. M. BRowN. 1981. Butterflies of the Rocky Mountain states. Univ. 
Oklahoma Press, Norman. 442 pp. 

GREY, L. P., A. H. MoECK, & W. H. Evans. 1963. Notes on overlapping subspecies II. 
Speyeria atlantis in the Black Hills. J. Lepid. Soc. 17:129-147. 

McCaBE, T.L. 1981. The dakota skipper, Hesperia dacotae (Skinner): range and biology, 
with special reference to North Dakota. J. Lepid. Soc. 35:179-198. 

MASTERS, J. H. 1975. Butterflies observed in Sieche Hollow State Park, South Dakota. 
J. Lepid. Soc. 29:126-127. 

MILLER, L. D. & F.M. BROWN. 1981. A catalogue/checklist of the butterflies of America 
morthlof Mexico. Mer! Lepid. Soc. 2:1—280. 

NoRDIN, J. S. 1967. Hesperiidae records for South Dakota. Newsl. Assoc. Minn. Entomol. 
1:90-92. 

. 1968. 1966 Lycaenidae records for South Dakota. Bull. Assoc. Minn. Entomol. 
2:60-61. 

ScoTT, J. A. & R. E. STANFORD. 1981. Geographic variation and ecology of Hesperia 
leonardus (Hesperiidae). J. Res. Lepid. 20:18-35. 


VOLUME 48, NUMBER 3 247 


STANFORD, R. E. & P. A. OPLER. 1993. Atlas of western USA butterflies including 
adjacent parts of Canada and Mexico. Published by the authors. 

TRUMAN, P. C. 1896. Lepidoptera in South Dakota. Entomol. News 7:298-299. 

1897. Lepidoptera in South Dakota. Entomol. News 8:27-29. 


Received for publication 10 April 1993; revised and accepted 15 February 1994. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 248-257 


BIOLOGY OF ANAEA RYPHEA (NYMPHALIDAE) IN 
CAMPINAS, BRAZIL 


ASTRID CALDAS 


Departamento de Biologia Animal e Vegetal—IB, 
Universidade do Estado do Rio de Janeiro, 
Rio de Janeiro, RJ 20550-0138, Brazil . 


ABSTRACT. Anaea ryphea uses Croton floribundus (Euphorbiacae) as its main larval 
food plant at Campinas, Brazil. Weekly censuses of the immature stages of A. ryphea 
were conducted from September 1988 to August 1989. Adults and larvae were found 
only from December through May. Females usually laid one egg per leaf and exhibited 
no plant height preference. Within individuals plants, most eggs were laid on the inter- 
mediate leaves; they were rare on the lowest and absent on the apical, new leaves. The 
complete life cycle in the field lasts 50 to 60 days. The pattern of development in A. 
ryphea is similar to that described for 5 other species of Anaea. The early stages resemble 
closely those described for A. eurypyle, which also uses a species of Croton as its larval 
food plant in El Salvador. 


Additional key words: Hypna clytemnestra, life cycle, Memphis, Croton, Euphor- 
biaceae. 


The genus Anaea Hubner includes most of the Neotropical Char- 
axinae, although use of the generic name varies considerably among 
authors. Comstock (1961) assigned to Anaea the species now considered 
members of the Anaea troglodyta group. He used Memphis Hubner, 
formerly described as a generic name, as a subgenus for most of the 
other species of Anaea, including the blue species and A. ryphea (Cra- 
mer). Rydon (1971) subdivided the group further, describing Foun- 
tainea, into which he transferred ryphea. Although the most recent 
treatment of the group (Descimon 1986) recognizes Anaea as distinct 
from Fountainea, other authors (e.g., DeVries 1987) include Fountai- 
nea species in Memphis. According to Hemming (1967), Anaea has 
priority, leaving Memphis a synonym. 

During the last ten years, Anaea andria Scudder has been the subject 
of several studies, and its biology is now well known. It uses Croton 
capitatus (Euphorbiaceae) as its primary larval food plant in North 
America (Riley 1981, 1988, 1989). The life cycles of other species of 
Anaea have been reported by Muyshondt (1973, 1974a, 1974b, 1975a, 
1975b); however, the early stages of A. ryphea have remained unknown. 
DeVries (1987) reported Anaea ryphea from Mexico to the Amazon 
Basin, but it has become evident that its distribution is wider than 
previously thought. In southern Brazil it uses Croton floribundus Spreng 
(Euphorbiaceae) as its primary larval food plant (Caldas 1991). 

Anaea ryphea is sexually dimorphic: males are smaller and usually 
lack the tail-like expansion of the hindwing (Figs. 1 & 2). Males are 
bright orange, red, and purple, with blue distal bars on the forewing cells. 


VOLUME 48, NUMBER 3 ; | 249 


Fics. 1-2. Ventral (left) and dorsal (right) views of adult Anaea ryphea. 1, male. 2, 
female. 


Females always have the hindwing expansion (“‘tail’’), dark brown bars 
on the forewing cells, and yellow and orange as the dominant colors 
on both wings. The species is found in tropical and subtropical forests, 
where it can be seen in rapid flight or perched on urine, feces, and 
rotten fruits. Herein I describe the life cycle of A. ryphea from south- 
eastern Brazil. 


STUDY SITE AND METHODS 


This study was conducted in the Santa Genebra Reserve, Campinas, 
Sao Paulo, Brazil (22°54’S, 47°05’W, elevation 650 m). The reserve 
includes 2,517,759 m2 of disturbed subtropical semi-deciduous forest. 
Mean monthly temperature varies from 18°C to 29°C, with daily fluc- 
tuations of as much as 20°C from July to September. The dry season 
lasts from May through September. The rainy season may start with 
light rainfall or with heavy “summer storms.” Frosts are rare, but may 
occur from July to September. Although the study was conducted from 
September 1988 through April 1991, the data presented here refer only 
to the first wet season, from December 1988 to May 1989. 

I visited the reserve once a week from September 1988 through 
January 1989, then twice a week until mid-February. From then on, 
visits were more frequent, increasing to every other day until late April 
1989, when they became weekly again. 

Croton floribundus is a lactescent plant that can be either a small 
shrub or a tall tree, depending on its age and environmental conditions. 
The smallest plant found was only 20 cm tall, while the largest was 
more than 4 m. The leaves are alternate, stipulate, and grayish green. 


250 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Leaf size depends on plant height: 10 cm long on plants up to 2 m tall 
and 20 cm long on taller plants. The inflorescences are monoecious, 
with male flowers terminal. At the study site, flowers were seen during 
December. Other species of Croton are recorded as food plants for 
species of Nymphalidae (DeVries 1987) and Riodinidae (DeVries 1988). 
Croton floribundus is common in the Santa Genebra Reserve, in both 
sunny and shaded areas, as solitary individuals or in patches. It grows 
mostly along the edges of trails and is rare in the middle of the forest. 
The soil along the trails is mostly clay, and during the dry season the 
plants seem to resist desiccation. From May on, plants begin to dry out, 
sometimes being reduced to no more than a woody stem that puts out 
new leaves the next wet season. During the rainy season, the soil main- 
tains a high water content. Along the reserve’s central road (1,160 m 
long), I found 186 plants. All were mapped and marked for use in this 
study. 

During each visit I checked all leaves for larvae on the 186 marked 
plants. Eggs were difficult to detect, so exact numbers were not known 
until the following season. Twenty eggs were collected and taken into 
the laboratory. These eggs were the source of hatching time observa- 
tions. In the field, each leaf with a larva was individually numbered 
because larvae stay on the same leaf most of the time. All larvae were 
measured and identified as to instar. Their height from the ground and 
the general aspect of the leaf were noted. With this methodology I was 
able to (1) follow the growth of every individual, (2) determine the 
duration of each instar in the field, and (3) assess mortality for each 
instar. 


RESULTS AND DISCUSSION 
Oviposition 


Anaea ryphea eggs are laid singly and usually one per leaf, in the 
central third of the under surface. Rarely, two or three eggs are laid. 
A similar oviposition pattern was described by Ramos (1984) for Anaea 
troglodyta borinquenalis and by Muyshondt (1975b) for Anaea (Mem- 
phis) pithyusa. Females fly around the plant, inspecting leaves quickly 
before ovipositing. I could not tell whether eggs laid on the same leaf 
came from the same female. Placing one egg per leaf could eliminate 
competition during the early instars, when larvae have restricted move- 
ment. In Colon Province, Republic of Panama, 1991, while studying 
the same species, I saw females placing more than one egg on the same 
leaf of Croton billbergianus, although not sequentially. In each of the 
three cases observed, the female flew from one leaf to another, ovi- 


VOLUME 48, NUMBER 3 | 251 


positing on acceptable leaves, eventually returning to oviposit on a leaf 
she already had used. 

There was no plant height preference for oviposition among the 186 
marked plants. All instars were found between 15 cm and 2 m above 
the ground, on short or tall plants. I did not include plants taller than 
2 m in this study owing to the difficulty in reaching high leaves, but 
even on those, I could see active larvae when they were present. Within 
the same plant, though, larvae were found mostly on intermediate 
leaves, never on the apical ones, and rarely on the lower (older) ones. 
This distribution seems to be related to the age of the leaves, not to 
their height from the ground. 


Development and Behavior 


After eclosion larvae eat the egg shell and move toward the apex of 
the leaf where they begin to feed on both sides of the midvein, which 
when exposed becomes their resting place. This behavior, common in 
Nymphalidae, also is considered common for other Anaea species (Muy- 
shondt 1974a). Ramos (1984) reported that larvae of A. troglodyta 
borinquenalis build a perching place from the tip instead of exposing 
the midvein first. When not feeding, A. ryphea larvae remain on this 
vein facing the apex, grasping the vein with both thoracic and abdom- 
inal legs. It is rare to see a larva start feeding on another part of the 
leaf, but when this happens, a lateral vein is chosen and used in the 
same manner. In either case, an individual may add fecula (‘‘frass 
chain’’) to the vein, perhaps to strengthen it. Fecula is added mostly 
when the vein is thin, whether it is a lateral or central one. It also may 
be used to extend the vein beyond its natural length. 

From the third instar on, A. ryphea larvae may roll the leaf to form 
a tubular shelter, holding it closed with silk, although this behavior 
usually is observed from fourth instar on. They begin by laying silk 
over the entire area of leaf that is to be rolled. As they add to and 
strengthen the threads, the leaf begins to curve and roll itself into a 
tube. Normally, the apical portion of the leaf is used to construct the 
shelter. If the leaf is damaged by some external event such as bird 
attack or herbivores, the larva may move to another leaf. Change of 
leaf does not necessarily follow any special rule, and I have observed 
larvae moving to new and old leaves, above or below the original one. 
The tube is open at both ends, but the opening that faces the base of 
the leaf is the larger of the two and is used by the larvae as the entrance 
and exit. The smaller opening is used to dispose of fecula. In order to 
enter the tube, the larva approaches close to opening, then makes a 
u-turn and backs into the shelter. 

I did not measure leaf consumption, but observations show that larvae 


Joe JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


usually feed on the same leaf through the third instar, reducing it to 
one-half or one-third its original size. From third instar on, feeding 
increases noticeably. Larvae can move to another leaf and build a new 
tube if the first leaf is consumed before pupation, but that rarely was 
seen. More commonly they visit another leaf just to eat, then return to 
the old leaf where the shelter is kept (they never eat the roll). Feeding 
takes place mainly from dusk to dark, as do molting and pupation. 
When not eating, larvae remain on the midvein or inside the tube. 
During the last instar, the larva abandons the plant and searches for a 
suitable place to pupate. 


Early Stages 


Egg (Fig. 3). Approximately 1 mm in diameter (n = 10). Spherical, with small depression 
on top (micropyle end), greenish white, with no conspicuous sculpturing. Duration 5-6 
days. 

First Instar (Fig. 5). Length 1.5-8.0 mm (n = 1161). Head brown with dark brown 
spots, lighter near foramen; epicranial suture complete, well marked. Body eruciform, 
hazel, darker posteriorly. Duration approximately 6 days (n = 278). 

Second Instar (Fig. 4). Length 5-14 mm (n = 598). Head light brown with three dark 
stripes on each side; frons with central dark stripe; pair of small scoli (each with two 
setae) on vertex, scattered chalazae restricted to sides of head. Body pale greenish brown 
with pale spots throughout, slightly enlarged on second abdominal segment. Duration 
approximately 6 days (n = 155). 

Third Instar (Fig. 6). Length 8-20 mm (n = 308). Head similar to that of second instar, 
but with a series of dark spots between the three dark stripes of the epicranium, also 
some dark dots around the foramen; scoli larger than on second instar, with more chalazae. 
Body green, except second, fifth, and seventh abdominal tergi, which are brown; second 
abdominal segment enlarged, forming a small dorsal hump. Duration approximately 7 
days (n = 105). 

Fourth Instar (Fig. 7). Length 14-24 mm (n = 301). Head similar to third instar, with 
dark stripes more pronounced, scoli resembling antlers, and chalazae all over head. Body 
brownish green, with dark brown subdorsal stripes following second and third thoracic 
segments, turning dorsal and joining to form a broad dorsal band across the first and 
second abdominal segments; fourth through tenth abdominal segments also with dark 
brown subdorsal stripe; third abdominal segment entirely green. Hump on second ab- 
dominal segment more conspicuous than in previous instar. Duration approximately 7 
days (n = 83). 

Fifth Instar (Fig. 8). Length 20-28 mm (n = 273). Remarkably distinct from previous 
instars. Head green; scoli large, Y-shaped; chalazae present in greater density. Body bright 
green, with black dots on dorsal and lateral surfaces of all segments; second, fifth, and 
seventh abdominal segments with reddish hazel subdorsal stripes. Duration 8-15 days (n 
= 45). 

Pre-pupa (Fig. 9). Green, curled horizontally rather than vertically as in other nym- 
phalids, and appressed to its support. Duration 2 days (n = 12). 

Pupa (Fig. 10). Green, naked; dark cremaster attached to white silken pad. Duration 
approximately 12 days (n = 12). 


The duration of the early stages in the field requires 50-60 days, 
accounting for individual variation. The pattern of development of 
Anaea ryphea is similar to that described for Anaea (Zaretis) itys on 


VOLUME 48, NUMBER 3 . DSI) 


Fics. 3-6. Early stages of Anaea ryphea. 3, Egg on Croton billbergianus leaf. 4, 
Second instar larva with frass chain. 5, First instar. 6, Third instar larva. 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


VOLUME 48, NUMBER 3 255 


Casearia nitida (L.) Jacq. (Flacouidaceae) (Muyshondt 1973); for Anaea 
(Consul ) fabius Doubleday on Piper tuberculatum Jacquin, P. auritum 
H.B.K., and P. umbellatum L. (Piperaceae) (Muyshondt 1974a); and 
for Anaea (Memphis) pithyusa on Croton reflexifolius H.B.K. and C. 
niveus Jacquin (Euphorbiaceae) (Muyshondt 1975b), all of which were 
reared in the laboratory from field collected eggs. The eggs of the four 
species (including A. ryphea) are similar, and instars show minor vari- 
ations. Duration of development is slightly shorter in A. (Zaretis) itys 
and A. (Memphis) pithyusa, and longer in A. (Consul) fabius. It is 
likely that the differences observed in the laboratory also would be 
present under field conditions. Anaea (Zaretis) itys differs from the 
others in that it does not build a leaf roll. Instead, it remains at the tip 
of the leaf until the pre-pupal stage. In all four species, the pre-pupae 
curl horizontally instead of vertically as in other nymphalids, and hang 
straight down only after pupation. Ramos (1984) described the pre- 
pupa of A. troglodyta borquenalis as different from these four species— 
it is the only one that curls vertically. 

In addition to the species discussed above, the early stages of Anaea 
eurypyle confusa Hall (Muyshondt 1974b) also are similar to those of 
A. ryphea. Both use species of Croton, and the eggs and larvae are 
similar. There are slight differences in head sculpturing and body color 
of the larvae, and in color and shape of the adult’s wings. 

The life history information presented above on A. ryphea was col- 
lected during extensive field observations. The animals in my study 
were allowed to remain under natural conditions in contrast to the 
laboratory conditions of the other studies cited above. This is one of 
the reasons for the dramatic decrease in numbers of observations from 
one instar to the next, which more accurately reflects the fate of natural 
populations in the field. Only the pre-pupa and pupa stages had to be 
observed in the laboratory because fifth instar larvae would have been 
lost when they left their plants to pupate. No pupae were ever found 
on or near C. floribundus in the field. 

Variations in rearing conditions can affect the ultimate form of the 
adult in species with seasonal forms. This was studied by Riley (1981, 
1989) in A. andria. So far, no seasonal variation has been observed in 
A. ryphea in the field, although males exhibit considerable variation in 
the color and shape of the wings. 


— 


Fics. 7-10. Early stages of Anaea ryphea. 7, Fourth instar larva. 8, Fifth instar larva. 
9, Pre-pupa. 10, Pupa. 


256 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Herbivory and Associated Fauna 


Anaea ryphea larvae seldom were seen eating the lowest leaves (old 
and damaged), and they never fed on apical leaves. Other herbivores 
also were observed on the intermediate portions of the plants. One 
species of cricket (Orthoptera: Gryllidae) ate patches from the leaves 
giving them the aspect of lace; one species of beetle (Coleoptera: Chrys- 
omelidae) was observed eating the petiole; and one species of true bug 
(Hemiptera) appeared to be feeding on the petiole. Larvae of Hypna 
clytemnestra Cramer (Nymphalidae) are common on Croton floribun- 
dus, frequently outcompeting A. ryphea with their ability to defoliate 
small plants. There also is a species of aphid (Homoptera: Aphidae) 
that uses C. floribundus. Individuals aggregate on the undersides of 
leaves, and as a result, the upperside becomes dotted with white spots. 

A leaf-mining microlepidopteran was taken to the laboratory for 
rearing, but all individuals died before pupation; hence, identification 
was not possible. The mine follows the edge of the leaf and when that 
area is used, the larva turns toward the central part of the leaf following 
the central vein, never crossing it. Pheidole ants (Hymenoptera: For- 
micidae) wee seen predating upon eggs of A. ryphea (n = 10), but were 
never observed interacting with the larvae. 

In addition to those species mentioned above, a variety of other insects 
and spiders were seen regularly on C. floribundus, but not feeding on 
it. Some individuals of C. floribundus produce globs of a translucent 
gum, usually close to the apical leaves, that perhaps serve as a me- 
chanical defense against some herbivores. 

The practice of leaving the midvein and secondary veins intact, as 
exhibited by A. ryphea, appears to represent a feeding strategy de- 
scribed for other species of Lepidoptera (Compton 1987), and differs 
from the strategy of those species that cut the midvein to prevent toxins 
from getting to the portion of the leaf to be consumed. The former 
strategy allows the larvae of A. ryphea to avoid contact with the latex 
that C. floribundus produces, which suggests that they are incapable 
of detoxifying or storing high concentrations of this substance. Hypna 
clytemnestra and the leaf-mining microlepidoptera reported from C. 
floribundus also avoid the midvein, although the former may eat sec- 
ondary veins. This type of behavior has been interpreted as an important 
step toward a disguise for these larvae (Hingston 1932) which add pieces 
of leaves and fecula to the midvein to improve the resemblance of the 
substrate to their body (Fig. 4). Such a behavior may reduce the level 
of predation. 


VOLUME 48, NUMBER 3 , Dol 


ACKNOWLEDGMENTS 


This work was supported by grants from FAEP/UNICAMP and CAPES, Brazil. I thank 
Woodruff W. Benson for teaching me the methodology and for his support of my ideas 
during the project; Keith S. Brown, Jr., for invaluable advice on Lepidoptera; and the 
Smithsonian Tropical Research Institute, Panama, for the use of facilities during the 
preparation of this manuscript and studies of local populations of A. ryphea. I am especially 
indebted to Carl C. Hansen for the wonderful photographs and to Annette Aiello, who 
exhaustively revised the manuscript with me. 


LITERATURE CITED 


CALDAS, A. 1991. A population of Anaea ryphea (Nymphalidae) and its larval foodplant 
at Campinas, Brazil. J. Lepid. Soc. 45:68. 

COMPTON, S. G. 1987. Aganais speciosa and Danaus chrysippus (Lepidoptera) sabotage 
the latex defenses of their host plants. Ecol. Entomol. 12:115-118. 

Comstock, W. P. 1961. Butterflies of the American tropics. The genus Anaea, Lepi- 
doptera, Nymphalidae. American Museum of Natural History, New York. 214 pp. 

DESCIMON, H. 1986. The evolution of coloration in Neotropical Charaxinae: Adaptive 
strategies and cladogenesis (Lepidoptera, Rhopalocera). Bull. Soc. Zool. Fr. 111:261- 
296. 

DEVRIES, P. J. 1987. The butterflies of Costa Rica and their natural history. Princeton 
University Press, New Jersey. 327 pp. 

1988. The larval ant organs of Thisbe irenea (Lepidoptera: Riodinidae) and 
their effects upon attending ants. Zool. J. Linn. Soc. 94:379-393. 

HEMMING, F. 1967. The generic names of the butterflies and their type-species (Lep- 
idoptera: Rhopalocera). Bull. Brit. Mus. (Nat. Hist.) Entomol. Suppl. 9. 

HINGSTON, R. W. G. 1932. A naturalist in the Guiana forest. Longmans, Green & Co., 
New York. 

MUYSHONDT, A. 1973. Notes on the life cycle and natural history of butterflies of El 
Salvador. II. Anaea (Zaretis) itys (Nymphalidae). J. Lepid. Soc. 27:294-302. 

. 1974a. Notes on the life cycle and natural history of butterflies of El Salvador. 

III. Anaea (Consul ) fabius (Nymphalidae). J. Lepid. Soc. 28:81-89. 

1974b. Notes on the life cycle and natural history of butterflies of El Salvador. 

III. Anaea (Memphis) eurypyle confusa (Nymphalidae). J. Lepid. Soc. 28:306-314. 

1975a. Notes on the life cycle and natural history of butterflies of El Salvador. 

III. Anaea (Memphis) morvus boisduvali (Nymphalidae). J. Lepid. Soc. 29:32-39. 

1975b. Notes on the life cycle and natural history of butterflies of El Salvador. 
Ill. Anaea (Memphis) pithyusa (Nymphalidae). J. Lepid. Soc. 29:168-176. 

RAMOS, S. J. 1984. Notes on the life cycle and biology of Anaea troglodyta borinquenalis 
(Lepidoptera: Apaturidae). Carib. J. Sci. 20:19-24. 

RILEY, T. J. 1981. Effect of long and short day photoperiod on the seasonal dimorphism 
of Anaea andria (Nymphalidae) from central Missouri. J. Lepid. Soc. 34:330-337. 

1988. Effect of larval photoperiod on mating and reproductive diapause in 

seasonal forms of Anaea andria (Nymphalidae). J. Lepid. Soc. 42:263-268. 

. 1989. Effect of larval photoperiod on incidence of adult seasonal forms in Anaea 
andria (Lepidoptera: Nymphalidae). J. Kansas Entomol. Soc. 61:224—227. 

RYDON, A. H. B. 1971. The systematics of the Charaxidae (Lepidoptera: Nymphalidae). 
Entomol. Rec. J. Var. 83:339-341. 


Received for publication 25 June 1998; revised and accepted 12 March 1994. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 258-263 


METHODS FOR SEXING LEPIDOPTERA LARVAE USING 
EXTERNAL MORPHOLOGY 


DEssIE L. A. UNDERWOOD 


Section of Evolution and Ecology, Center for Population Biology, 
University of California, Davis, California 95616, USA 


Abstract. This paper describes a method for determining the sex of larval Lepidoptera 
using external morphology. This method, previously developed by others but published 
in journals with limited distribution, uses sex-specific pits located ventrally on the 8th 
and 9th abdominal segments. This paper also describes a new technique for clearing and 
staining newly hatched larvae so that these sex-specific pits can be identified readily. 
Finally, this paper extends the applicability of this method to four other species in three 
butterfly families (Papilionidae, Nymphalidae, and Pieridae). 


Additional key words: sex determination, larval morphology. 


The ability to sex live Lepidoptera larvae would allow for novel 
experiments on sex-specific differences in behavior, physiology, and 
development, as well as hormonal and metabolite assays, to name a 
few. Determination of sex ratio and sex-specific mortality typically has 
been limited to those larval stages where sex can be determined by 
dissection. Yet in most species gonad differentiation does not occur until 
at least the third instar, and in some species such as Papilio zelicaon 
Lucas (Papilionidae), even last instar larvae are impossible to sex by 
examining the gonads. Moreover, sexing by dissection necessarily in- 
volves sacrificing the animals—thereby precluding any further study 
of these individuals. 

This paper describes a method of wide application for sexing Lep- 
idoptera larvae using external morphology. It was developed by others 
but published in journals with limited distribution (Joseph & Karnavar 
1991, Lavenseau 1982, Muraleedharan & Muraleedharan 1989, Stehr 
& Cook 1968) and seems to be little known or used. Aside from Pieris 
rapae L. (Pieridae), this method has been tested previously only in the 
moth families Psychidae, Pyralidae, Notodonidae, Lymantridae, Noc- 
tuidae, Arctiidae, and Attacidae (Stehr & Cook 1968, Lavenseau 1982, 
Muraleedharan & Muraleedharan 1989, Joseph & Karnavar 1991). 

Larvae can be sexed live once they are large enough, typically at 
least 5 mm in length, to allow one to clearly see the last four abdominal 
segments under a dissecting microscope. All instars of preserved larvae 
can be sexed. This paper extends the applicability of this method to 
four other species within three butterfly families. It also presents a new 
technique for clearing and staining very young preserved larvae, in- 
cluding first instars, so that this method can be used to determine their 
SEX. 


VOLUME 48, NUMBER 3 | 259 


MATERIALS AND METHODS 


Fifth instar larvae of Eucheira socialis Westwood (Pieridae), Battus 
philenor Linnaeus (Papilionidae), Junonia coenia Hubner (Nymphal- 
idae), and Papilio zelicaon were sexed readily by examining the 8th 
and 9th abdominal segments using a Wild dissecting microscope. I first 
sexed individuals using external morphology, then dissected and sexed 
the same individuals by examining their gonads. I examined at least 
10 larvae, five of each sex, from each of the four species of butterflies. 
All sex determinations were confirmed by dissection for all species 
except Papilio zelicaon where gonadal differentiation was insufficient 
in fifth instar larvae to determine sex. For this species, I sexed individuals 
by external morphology, then reared them out and sexed them as adults. 

First instar larvae of E. socialis preserved in a 1:3 part mixture of 
glacial acetic acid and methanol were placed in a petri dish with 
glycerin and acetic acid (1:4). Petri dishes with larvae were put into 
larger covered petri dishes to prevent excessive evaporation, then placed 
in a oven at 60°C for two hours. Petri dishes were removed from the 
oven and left at room temperature for 22 hours. The front half of each 
larva was removed with forceps. The remaining rear half was placed 
in Hoyer’s medium, prepared using the protocol in Ashburner (1989), 
and then incubated in an oven at 37°C for 18 hours. Slides were prepared 
with Hoyer’s as the mounting medium with larvae oriented ventral 
surface up. I viewed slides under epi-illumination using a plan 16 
objective, 50W mercury bulb, cross polars, a heat filter, and a green 
interference filter. Slides prepared in this manner should be useful for 
several weeks, but eventually will degrade. 


RESULTS 


Sex determined by external morphology was confirmed by dissection 
or sexing adults in all but one larva. The only erroneously sexed larva, 
a P. zelicaon, belonged to the species most difficult to sex because larvae 
were light in color and the pits lacked contrasting coloration. 

Sex-specific pits were clearly visible on the ventral surface of larvae 
(Figs. 1 and 2). Males were characterized by having one central pit on 
the 9th abdominal segment. Females had four pits, two located on each 
of the 8th and 9th abdominal segments. 

For larvae requiring clearing and staining, the time necessary for 
the initial treatment of glycerin and acetic acid and for the Hoyer’s 
treatment probably will depend upon the particular characteristics of 
the larvae to be examined. Some species may require more complete 
clearing and longer incubation times at perhaps greater temperatures, 
while others may not require clearing at all. The best preparations of 


260 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


we era: 


Fic. 1. First instar Eucheira socialis (Pieridae). (A) female (160) with four pits, and 
(B) male (160) with one pit; pits indicated by arrows. See text for details on microscope 
settings. Fifth instar E. socialis. (C) female (16x) with four pits, and (D) male (16x) with 
one pit. All animals in the photos are preserved and, except (A), are oriented with the 
posterior end downward. 


VOLUME 48, NUMBER 3 261 


ae 


FIG. Papilio zelicaon (Papilionidae) (A) female (20x), (B) male (16x); Junonia 
coenia 7 (C) female (18x), (D) male (20x). All animals in the photos are 
preserved and oriented with the posterior end downward. 


262 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


first instar E. socialis were those with partial clearing; second instars 
could be sexed readily without clearing but needed the additional mag- 
nification of the compound microscope and epi-illumination. 


DISCUSSION 


Depending upon age and/or species, it was sometimes easier to iden- 
tify one sex or the other. In E. socialis, female second to fourth instars 
were readily identified because the four pits were surrounded by lighter 
colored halos, while the male pits were relatively difficult to discern. 
Fifth and sixth instar male E. socialis were quickly identified because 
the pit typically was darkened while the female pits were less distinctive. 
In contrast, female J. coenia were much more distinctive than the males 
even though only one pair of pits in the female was predictably dis- 
cernable. Female P. zelicaon also were easier to sex than males, but 
both were sometimes difficult because these larvae are very light in 
color, and the pits lacked contrasting coloration. 

Lavenseau (1982) sexed first instars by searching for abdominal pits 
on larvae using scanning electron microscopy. This technique requires 
relatively elaborate protocol for preparing larvae. The technique for 
clearing and staining first instar larvae described here requires only a 
microscope capable of epi-illumination. The procedure used here for 
clearing larvae using Hoyer’s medium is used in our introductory em- 
bryology course at the University of California at Davis to study cu- 
ticular mutations in Drosophila embryos; hence, it is simple enough to 
be done successfully by the inexperienced. 

Two other techniques have been published to sex larvae with un- 
differentiated gonads which do not use sex-specific abdominal pits. The 
nuclei of many species of animals are known to contain darkly staining 
bodies which correlate with the presence of the Y chromosome (Smith 
1945a, 1945b, Ennis 1976, Traut & Scholz 1978, Clarke 1984). Clarke 
(1984) identified the sex of living larvae by removing a proleg and 
scraping enough tissue for preparations which stain heteropycnotic bod- 
ies. However, these heteropycnotic bodies are not always evident in 
females in some species of Lepidoptera, or rarely, they are seen in both 
sexes (Traut & Mosbacher 1968, Ennis 1976, Traut 1976). 

Seiler (1964), in a study on intersexes in a moth, Solenobia triquetrella 
F. R. (Psychidae), sexed larvae with undifferentiated gonads by follow- 
ing the gonoducts. In females, the gonoducts terminate at the 7th ab- 
dominal segment, and in males, they terminate at the 9th abdominal 
segment where they connect to the ectodermal anlage of the male 
copulation organ (the Organ of Herold). Newly hatched larvae were 
sexed by the presence or absence of the Organ of Herold. However, 


VOLUME 48, NUMBER 3 263 


this method cannot be used on living larvae no matter how mature, 
and I found the dissections tedious and difficult to perform. 


ACKNOWLEDGMENTS 


I thank Mark Camara and Sherri Graves for donating live larvae. Adam Porter lent 
me Stehr and Cook’s publication which first introduced me to this method. Peter Arm- 
strong made possible the photos taken through the epi-illuminating microscope and Judy 
Nelson provided assistance in photographing the fifth instars. Jeanette Natzle suggested 
using the clearing method for the first instars and she provided the Hoyer’s medium. 


LITERATURE CITED 


ASHBURNER, M. 1989. Drosophila—a laboratory manual. Cold Spring Harbor Labo- 
ratory Press, Cold Spring. 434 pp. 

CLARKE, C. 1984. Upsets in the sex-ratio of some Lepidoptera, pp. 255-258. In Vane- 
Wright, R. & P. R. Ackery (eds.), The biology of butterflies. Academic Press, New 
York. 

ENNIS, T. J. 1976. Sex chromatin and chromosome numbers in Lepidoptera. Canad. J. 
Genet. Cytology 18:119-130. 

JOSEPH, Fr. T. M. & G. K. KARNAVAR. 1991. Identification of sex in the larval and 
pupal stages of Ailanthus defoliator, Eligma narcissus indica Roth. (Lepidoptera: 
Noctuidae). Entomon 16:331-333. 

LAVENSEAU, L. 1982. Determination of sex of caterpillars without dissection. Int. J. 
Insect Morph. Embry. 11:359-362. 

MURALEEDHARAN, A. & D. MURALEEDHARAN. 1989. Biology and morphometrics of 
castor semilooper, Achoea janata Linn. (Lepidoptera: Noctuidae). Uttar Pradesh J. 
Zool. 9:48-55. 

SEILER, J. 1964. Sexuality as developmental process. Proc. Int. Con. Genet. 11:199-207. 

SMITH, S. G. 1945a. The diagnosis of sex by means of heteropycnosis. Sci. Ag. 25:566- 
aml: 

1945b. Heteropycnosis as a means of diagnosing sex. J. Heredity 36:194—-196. 

STEHR, F. W. & E. F. CooK. 1968. A revision of the genus Malacosoma Hibner in 
North America (Lepidoptera: Lasiocampidae): Systematics, biology, immatures, and 
parasites. U.S. Natl. Mus. Bull. 276:1-321. 

TRAUT, W. 1976. Pachytene mapping in the female silkworm, Bombyx mori L. (Lep- 
idoptera). Chromosoma 58:275-284. 

TRAUT, W. & G. C. MOSBACHER. 1968. Geschlechtschromatin bei Lepidoptera. Chro- 
mosoma 25:343-356. 

TRAUT, W. & D. SCHOLZ. 1978. Structure, replication and transcriptional activity of 
the sex-specific heterochromatin in a moth. Exp. Cell Res. 113:85-94. 


Received for publication 18 January 1994; revised and acceped 19 March 1994. 


GENERAL NOTES 


Journal of the Lepidopterists’ Society 
48(3), 1994, 264-265 


SOMATIC MOSAICISM IN MANIOLA JURTINA 
(NYMPHALIDAE: SATYRINAE) 


Additional key words: bilateral size asymmetry. 


Spontaneously appearing perfectly or nearly perfectly bilaterally differentiated, but 
non-gynandrous, individuals are known chiefly in the silkworm (Bombyx mori L., Bom- 
bycidae), fruit fly (Drosophila melanogaster L., Drosophilidae), and honeybee (Apis 
mellifera L., Apidae), and in some domesticated vertebrates. Selected examples from 
butterflies include female Papilio polytes L. (Papilionidae) with one side of the sex-limited 
f. “romulus’ Cramer and the opposite side of male-like f. “cyrus” Fabricius (Ollenbach 
1930); half black/half yellow morph females of Papilio glaucus L. (Papilionidae) (for a 
review see Clarke & Clarke 1983, Scriber & Evans 1988); and male Apatura ilia (Den. 
& Schiff.) (Nymphalidae) with one side of f. “silvia” Cabeau (Cockayne 1985). Apparently, 
asymmetries in butterflies involve mainly differences in wing pattern and coloration, and 
to a lesser extent, the shape and size of the wings. Some may represent cases of wing 
homoeosis (for a review see Sibatani 1980, 1983a, b). 

The teratology described herein concerns exclusively the size of body halves. The 
example was found in the field stock of two thousand specimens of the meadow brown, 
Maniola jurtina (L.) (Nymphalidae: Satyrinae), collected in July 1968 near Jarocin, Poland 
(51°58'N, 17°29’E), among lowland herb communities of the class Molinio-Arrhenather- 
etea. The aberrant specimen is a female (not a latent gynander) with the right part of 
the body distinctly smaller than the left, exhibiting ca. 8% linear difference (Fig. 1). The 
halving proceeds about the entire median line, equally affecting all structures of the right 
side of the head (e.g., palpus, antenna, compound eye), thorax (e.g., legs, wings with their 
scales and markings), and abdomen. For instance, the length of the left forewing (from 


Fic. 1. Bipartite mosaic female of M. jurtina. 


VOLUME 48, NUMBER 3 265 


base of costa to apex) is 23 mm, whereas that of the right is 21 mm (usual wing length 
in Polish populations is 23-25 mm). 

So far as I am aware, this represents the most obvious case of bilateral asymmetry in 
expression of insect body size, uncoupled with any other conspicuous asymmetric devi- 
ation, reported in the literature. The left-right difference in size is much more pronounced 
than that in bilateral gynandromorphs of M. jurtina figured by Barrington (1992), Al- 
brecht (1993), or Eitschberger (1993). 

As to their origin, “half-siders” in diploids are believed to result from point mutations, 
rearrangements, or malsegregation of chromosomes giving diversity at the first division 
of the zygote nucleus, or from double fertilization of a binucleate egg. In butterflies, 
mosaics thought to arise by polyspermy generally are gynandromorphic (preliminary 
separation of sister chromatids in meiosis) (Blanchard & Descimon 1988). Instead, non- 
disjunction of the homologues (i.e., XY/XO) recently has been proposed to be involved 
in appearance of the previously mentioned aberrations of P. glaucus (Clarke & Clarke 
1983, Scriber & Evans 1988). Comparative analysis of chromosomal DNA derived from 
the two halves of the aberrant specimens like that reported here may help in mapping 
loci that affect insect growth. 


LITERATURE CITED 


ALBRECHT, M. 1998. Ein gynandromorph von Maniola jurtina (Linnaeus, 1758) (Lep- 
idoptera, Satyridae). Atalanta 24:57-58, 318-319. 

BARRINGTON, R. D. G. 1992. Additional notes on aberrations of the meadow brown 
(Maniola jurtina L.) from North Dorset. Entomol. Rec. J. Var. 104:315-319. 

BLANCHARD, R & H. DESCIMON. 1988. Hybridization between two species of swallow- 
tails, meiosis mechanism, and the genesis of gynandromorphs. J. Lepid. Soc. 42:94— 
102. 

CLARKE, C. & F. M. M. CLARKE. 1983. Abnormalities of wing pattern in the eastern 
tiger swallowtail butterfly, Papilio glaucus. Syst. Entomol. 8:25-28. 

COCKAYNE, E. A. 19385. The origin of gynandromorphs in the Lepidoptera from bi- 
nucleate ova. Trans. Roy. Entomol. Soc. London 83:509-521. 

EITSCHBERGER, U. 1993. Ein weiterer gynandromorph von Maniola jurtina (Linnaeus, 
1758) (Lepidoptera, Satyridae). Atalanta 24:59-60, 318-319. 

OLLENBACH, O. C. 1930. Miscellaneous notes. XXIII.—A freak Papilio polytes romulus 
Cr. 2. J. Bombay Nat. Hist. Soc. 34:832-833. 

SCRIBER, J. M. & M. H. Evans. 1988. Bilateral gynandromorphs, sexual and/or color 
mosaics in the tiger swallowtail butterfly, Papilio glaucus (Lepidoptera: Papilionidae). 
J. Res. Lepid. 26:39-57. 

SIBATANI, A. 1980. Wing homoeosis in Lepidoptera: A survey. Devel. Biol. 79:1-18. 

1983a. A compilation of data on wing homoeosis in Lepidoptera. J. Res. Lepid. 

22:1-46. 

. 1983b. Compilation of data on wing homoeosis on Lepidoptera: Supplement 

I. J. Res. Lepid. 22:118-125. 


PRZEMYSLAW SZAFRANSKI, Boston University, Center for Advanced Biotechnology, 36 ~ 
Cummington Street, Boston, Massachusetts 02215, USA. 


Received for publication 1 May 1993; revised and accepted 28 November 1998. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 266 


HYMENOPTEROUS PARASITOIDS OF DISMORPHIA SPIO 
(PIERIDAE: DISMORPHIINAE) 


Additional key words: Trichospilus, Trichogramma, Puerto Rico. 


The following presents records of parasitoids of Dismorphia spio (Godart) reared from 
eggs and pupae collected during 1984-1986 in Barrio Guaonico, Utuado, Puerto Rico. 

During the study of the biology and natural history of D. spio (Torres Bauza 1991a), 
I collected five eggs parasitized by Trichogramma sp. (Trichogrammatidae). This rep- 
resents a parasitization rate of about 11% (n = 47) among the natural population. From 
each egg, up to six wasps emerged four to six days after collection. In Puerto Rico, 
Trichogramma sp. have been reported from eggs of Diatraea saccharalis (F.) (Pyralidae), 
Etiella zinckenella (Treitschke) (Pyralidae), Psara periusalis (Walker) (Pyralidae), Cal- 
podes ethlius (Cramer) (Hesperidae), and Eurema portoricensis (Dewitz) (Pieridae) (Wol- 
cott 1951, Torres Bauza 1991b). 

A pupa of D. spio collected on Inga vera Willd. (Mimosoideae) hosted Trichospilus 
diatraea Cherian & Margabandhu (Eulophidae). This represents a parasitization rate of 
about 1% (n = 84) among the natural population. Seventy wasps emerged from the pupa 
five days after collection. Trichospilus diatraea has been collected from Spodoptera 
frugiperda (Smith) (Noctuidae), Epimecis detexta (Walker) (Geometridae), Trichoplusia 
ni Huebner (Noctuidae), Pseudoplusia includens Walker (Noctuidae), Diaphania hyali- 
nata (L.) (Pyralidae), and Stenorrhages flegia (Cramer) (Pyralidae) in Florida, Trinidad, 
and Barbados (Bennet & Alam 1985, Bennet et al. 1987). This is the fist report of T. 
diatraea in Puerto Rico. 

Representatives of the reared parasitoids are deposited in the entomological collection 
of the Agricultural Experimental Station, University of Puerto Rico, Mayaguez Campus 
at Rio Piedras, Puerto Rico (lot number 90-10217). 

I thank Silverio Medina-Gaud for his assistance; M. E. Schauff and D. L. Vincent for 
parasite identifications; and two annonymous reviewers for constructive comments on the 
manuscript. 


LITERATURE CITED 


BENNET, F. D. & M. M. ALAM. 1985. An annotated check-list of the insects and terrestrial 
arthropods of Barbados. Carib. Agric. Res. Dev. Ins., Barbados. 81 pp. 

BENNET, F. D., H. GLENN, M. YASEEN & R. M. BARANOWSKI. 1987. Records of Tri- 
chospilus diatraea, an Asian parasite (Hymenoptera : Eulophidae) from the Caribbean 
and Florida. Florida Entomol. 70: 184-85. 

TORRES BAUZA, J. A. 199la. Biologia de Dismorphia spio (Godart) en Puerto Rico 
(Lepidoptera: Pieridae: Dismorphiinae). Carib. J. Sci. 27:35-45. 

. 1991b. Biologia de Eurema portoricensis (Dewitz) (Lepidoptera: Pieridae). 
Carib. J. Sci. 27:124—29. 

WOLCOTT, G. N. 1951. The insects of Puerto Rico. J. Agric. Univ. P. R. 32:537-748. 


JoRGE A. TORRES BAUZA, Department of Natural Sciences, La Montana Regional College 
(UPR), Call Box 2500, Utuado, Puerto Rico 00641-2500. Mailing address: Cabrera B-48, 
Utuado, Puerto Rico 00641. 


Received for publication 5 September 1993; revised and accepted 20 March 1994. 


Journal of the Lepidopterists’ Society 
48(3), 1994, 267-271 


OBITUARY 


ALBERT SCHWARTZ (1923-1992) 


The name Albert Schwartz is virtually synonomous with West Indian 
zoology. For nearly 40 years, Al devoted his life to the discovery, 
documentation, and description of butterflies, frogs, reptiles, birds, and 
mammals of the West Indian Islands. He probably was best known to 
lepidopterists for his book The Butterflies of Hispaniola and research 
on satyrid butterflies in the genus Calisto. Albert Schwartz died on 
Sunday, 18 October 1992 in a local hospital a few days after undergoing 
surgery for injuries sustained in a fall at his home in Miami, Florida. 

Schwartz was born on 13 September 1928 in Cincinnati, Ohio. He 
received his B.S. (Psychology) in 1944 from the University of Cincinnati, 
his M.S. (Zoology) in 1946 from the University of Miami, and his Ph.D. 
in 1952 from the University of Michigan. His master’s thesis topic was 
““Cestodes of sharks of the east coast of Florida” and doctoral dissertation 
topic was “Mammals of southern Florida and the upper Florida keys.” 
Al had a broad interest in zoology, but even as a graduate student, was 
drawn towards work on faunal surveys and descriptive studies. 

After completion of his doctorate, Al accepted a position in South 
Carolina as Curator of Vertebrate Zoology at the Charleston Museum. 
In 1956, he moved to Reading, Pennsylvannia and taught in the Biology 
Department of Albright College. Later he returned to Florida where 
he spent most of his professional career (1967-1988) teaching at Miami- 
Dade Community College. Because this was a non-research position, 
Al conducted his studies entirely on his own time. Even after retirement, 
Al jealously guarded his free time and carefully structured his life to 
maximize the time devoted to research on West Indian fauna and other 
favorite pursuits. Al was a gifted pianist, loved opera, and composed 
music as a hobby. A life-long bachelor, he enjoyed entertaining close 
friends but generally shunned large gatherings. 

Al’s West Indian field studies started in 1954 on Cuba, which re- 
mained his favorite island despite the inaccessibility to outside re- 
searchers in later years. In the late 1970s, having already accomplished 
many of his goals for surveying the herpetological fauna of the islands, 
Al began working on Lepidoptera and quickly realized that he could 
make significant contributions in this area of study. By the time of Al’s 
death, insect cabinets filled two bedrooms of his home, and shelving 
lined most of the wall space in living and dining rooms to house the 
oversized drawers with his butterfly collection. The collection includes 


268 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


c on / 


poh. : : E 


Fic. 1. Albert Schwartz in the early 1970s. Photo courtesy of M. H. Strahm. 


more than 28,500 specimens primarily from the West Indies and Flor- 
ida, with detailed locality data and documentation. 

The fauna of the islands captivated Al Schwartz and the scientific 
output which resulted was enormous. He published 280 papers (some 
5,100 pages) on West Indian zoology, including descriptions for three 
new species of bats, nearly two dozen species of butterflies, 80 species 
of amphibians and reptiles, and about 279 subspecies of frogs, lizards, 
and snakes. At the time of his death, he was collaborating on several 
papers dealing with West Indian mammals, amphibians and reptiles, 
and Lepidoptera (including a color atlas of Hispaniolan butterflies with 
Fernando Gonzalez). 

Students often accompanied Al in the field, and he also subsidized 
fieldwork for many of his collaborators. This became especially critical 
in his later years as chronic arthritis left Al increasingly incapacitated 


VOLUME 48, NUMBER 3 269 


and he had to rely on others to collect additional specimens and data. I 
came to know Al through my co-workers, Robert W. and Rose M. 
Henderson. Bob is a herpetologist and Rose an artist with the Milwaukee 
Public Museum. They both collaborated with Al on various projects 
and developed a warm friendship. While I never had the pleasure of 
meeting Al, I thoroughly enjoyed our phone conversations. Al had a 
wonderful sense of humor. He was a very open and direct person, and 
a great teacher, always encouraging and stimulating new ideas. Al 
Schwartz enriched the lives of numerous other lepidopterists, and his 
research legacy will serve as an inspiration for those studying the fauna 
of the West Indies for many years to come. 

As stipulated in his will, the Schwartz butterfly collection has been 
donated to the Milwaukee Public Museum by Michael H. Strahm, friend 
and former student of Al Schwartz, and executor of his estate. I owe 
special thanks to Bob Henderson for introducing me to Al, and for 
sharing fond remembrances and specific details about Al’s life for this 
article. 


SUSAN SULLIVAN BoRKIN, Invertebrate Zoology Section, Milwaukee Public Museum, 800 
West Wells Street, Milwaukee, Wisconsin 53233, USA. 


Received and accepted for publication 1 December 1998. 


LEPIDOPTERA PUBLICATIONS BY ALBERT SCHWARTZ 
1980 


1. (with W. W. Sommer) A new species of Atlantea (Nymphalidae) 
from Hispaniola, West Indies. Bull. Allyn Mus. 58:1-4. 

2. Deforestacion y Rhopalocera una comparacion en la recoleccion 
de mariposas entre Republica Dominicana y Haiti. Nat. Postal 30/ 
80: 1-8. 


1982 
3. (with C. J. Jimenez) The butterflies of Montserrat, West Indies. 
Bull. Allyn Mus. 66:1-18. 
4. Variation in Hispaniolan Greta diaphana (Ithomiidae). Bull. Allyn 
Mus. 69:1-10. 
1983 


5. A new Hispaniolan Calisto (Satyridae). Bull. Allyn Mus. 80:1-10. 
6. Haitian butterflies. Mus. Nac. Hist. Nat., Santo Domingo. 69 pp. 


270 


10. 


Jidhe 


2a. 


13. 


14. 


15. 


16. 


ee 


18. 


19. 


20. 


21. 


22. 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


. (with F. Gali). Myscelia antholia (Nymphalidae) in the Republica 


Dominicana. J. Lep. Soc. 37:164-165. 


. (with F. Gali). The second specimen of Epargyreus spanna (Hes- 


periidae). J. Lep. Soc. 37:170-171. 


. (with F. Gali). Battus zetides in the Reptblica Dominicana. J. Lep. 


Soc. 37:171-174. 


1984 


(with F. Gali). Five new species of Calisto (Satyridae) from His- 
paniola. Bull. Allyn Mus. 85:1-18. 


1985 


(with R. W. Wisor). Status of Calisto pulchella darlingtoni Clench 
(Lepidoptera: Satyridae). Florida Sci. 48:7-13. 

(with W. W. Sommer and F. Gali). Synapte mailitiosa [sic] (Lep- 
idoptera: Hesperiidae) on Hispaniola. Florida Sci. 48:13-17. 

A new subspecies of Calisto (Satyridae) from Hispaniola, West 
Indies. Bull. Allyn Mus. 93:1-5. 

(with J. Y. Miller). A new species of hairstreak (Lycaenidae) from 
Hispaniola. Bull. Allyn Mus. 99:1-6. 

(with J. C. Correa). The status of Calisto hysius batesi (Lepidoptera, 
Satyridae) with the description of a new species of Calisto from 
Hispaniola. Florida Sci. 49:11-18. 

(with W. W. Sommer). A new subspecies of Synapte malitiosa 
(Lepidoptera: Hesperiidae) from Hispaniola. Florida Sci. 49:18-22. 
(with F. Gali). Butterflies of the National Parks of Haiti. Mimeo., 
Florida State Mus. 19 pp. 


1987 


The taxonomic status of Pseudochrysops (Lycaenidae) on Puerto 
Rico. Bull. Allyn Mus. 110:1-5. 

(with F. L. Gonzales and R. M. Henderson). New records of but- 
terflies from the West Indies. J. Lep. Soc. 41:145-150. 

The butterflies of the lower Florida Keys. Contr. Biol. Geol. Mil- 
waukee Public Mus. 73:1-384. 


1988 


A new species of Calisto (Satyridae) from Hispaniola. Florida Sci. 
90:246-252. 
Butterflies of the Sierra Martin Garcia, Republica Dominicana. 
Carib. J. Sci. 23:418-431. 


VOLUME 48, NUMBER 3 DATEL 


23. 


24. 


20. 


26. 


27. 


28. 


29. 


30. 


ok 


(with F. L. Gonzalez). A new species of Calisto (Satyridae) from 
Hispaniola. Bull. Allyn Mus. 117:1-5. 


1989 


Rediscovery of Adelpha lapitha Hall, 1929 (Nymphalidae) on His- 
paniola, West Indies. Bull. Allyn Mus. 124:1-7. 
The butterflies of Hispaniola. Univ. Florida Press. xiv + 580 pp. 


1990 


(with R. W. Henderson). The butterflies of Barbuda, West Indies. 
Carib. J. Sci. 26:98-100. 


ISI 


(with F. L. Gonzalez and D. K. Wetherbee). A new species of 
Calisto (Lepidoptera: Satyridae) of the lyceia complex on Hispan- 
iola. Contr. Biol. Geol. Milwaukee Public Mus. 80:1-8. 

A new species of Calisto (Lepidoptera: Satyridae) from the Cor- 
dillera Central, Republica Dominicana. Contr. Biol. and Geol. Mil- 
waukee Public Mus. 81:1-3. 


1992 


(with K. Johnson). Two new butterflies (Lepidoptera: Lycaenidae) 
from Cuba. Carib. J. Sci. 28:149-157. 


In Press 


(with R. M. Henderson). The butterflies of St. Vincent, the Gren- 
adines, and Grenada, West Indies. Contr. Biol. Geol. Milwaukee 
Public Mus. 

(with F. L. Gonzalez). A color atlas of Hispaniolan butterflies. R & 
A Publ., England. 


Date of Issue (Vol. 48, No. 3): 17 August 1994 


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CONTENTS 


NECTAR SOURCE DIVERSITY AS AN INDICATOR OF HABITAT 
SUITABILITY. FOR THE ENDANGERED ~UNCOMPAHGRE 
FRITILLARY, BOLORIA ACROCNEMA (NYMPHALIDAE) Hugh 
B. Britten aa Byrn Riley 20 sees a 173 


HIGH ANDEAN PRONOPHILINI FROM VENEZUELA: Two NEW SPE- 
CIES OF DIAPHANOS (NYMPHALIDAE: SATYRINAE) Angel L. . 
Viloria Pit ee oe 180 


CONTINUED INTERSPECIFIC HYBRIDIZATION BETWEEN LIMENITIS 
(BASILARCHIA) ARTHEMIS ASTYANAX AND L. (B.) ARCHIPPUS 
IN THE SOUTHEASTERN U.S.(NYMPHALIDAE) Austin P. Platt 
and, James R. Maudsley: 3 2000 )) oot) ae 190 


FIELD OBSERVATIONS OF MATINGS BETWEEN FEMALE LIMENITIS 
ARCHIPPUS AND MALE L.. ARTHEMIS SUBSPECIES (NYMPHALIDAE) 


Charles'V., Covell) Jr" SiO a 199 
A TECHNIQUE FOR SETTING AND MOUNTING MICROLEPIDOP- 
TERA Jean-Francois Landry and Bernard Landry _. 205 
CHECKLIST OF SOUTH DAKOTA BUTTERFLIES (HESPERIOIDEA AND 
PAPILIONOIDEA) ) Gary) M. Marrone: 2... 228 
BIOLOGY OF ANAEA RYPHEA (NYMPHALIDAE) IN CAMPINAS, 
BRAZIL. Astrid Caldas, (2 ies) he hd i QAS 
METHODS FOR SEXING LEPIDOPTERA LARVAE USING EXTERNAL 
MORPHOLOGY /Dessie L. A. Underwood — 3. 1 ae 258 


GENERAL NOTES 


Somatic mosaicism in Maniola jurtina (Nymphalidae: Satyrinae) 


Przemyslaw;Saafranskt; 220 8ob) aN 264 
Hymenopterous. parasitoids of Dismorphia spio (Pieridae: Dismorphiinae) — 
jorge A:>Torres Bauzd .2 0000 CEs ee Se i ee 266 
OBITUARY 


Albert Schwartz (1923-1992) Susan Sullivan Borkirr ccc cece 267 


This paper meets the requirements of ANSI/NISO Z39.48-1992 (Permanence of Paper). 


NT Volume 48 3 1994 Number 4 


ISSN 0024-0966 


JOURNAL 


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‘LEPIDOPTERISTS’ SOCIETY 


Published quarterly by THE LEPIDOPTERISTS’ SOCIETY 


Publié par LA SOCIETE DES LEPIDOPTERISTES 
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- 1995 
on 


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19 December 1994 


THE LEPIDOPTERISTS’ SOCIETY 


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Cover illustration: The largest North American silverspot, the great spangled fritillary, 
Speyeria cybele (Fabricius) (Nymphalidae). Original drawing by Mark Etheridge, 9422 Fair- 
leigh Court, Burke, Virginia 22015. 


JOURNAL OF 


Tue LEpPIDOPTERISTS’ SOCIETY 


Volume 48 1994 Number 4 


Journal of the Lepidopterists’ Society 
48(4), 1994, 273-337 


GENITALIA AT THE GENERIC LEVEL: 
ATRYTONE RESTRICTED, ANATRYTONE RESURRECTED, 
NEW GENUS QUASIMELLANA—AND YES! WE HAVE 
NO MELLANAS (HESPERIIDAE) 


JOHN M. BURNS 


Department of Entomology, National Museum of Natural History, 
Smithsonian Institution, Washington, D.C. 20560, USA 


ABSTRACT. Some prior opinion notwithstanding, both male and female genitalia 
are exceedingly valuable in delimiting genera of skippers and in grouping species within 
genera. Even such small genitalic parts as the cornutus and juxta of the male can have 
value out of all proportion to their size. 

The New World genera Atrytone Scudder and Mellana Hayward are polyphyletic. 
Atrytone genitalically sorts out into (1) Atrytone, with the one species Atrytone arogos 
(Boisduval & Leconte) from the central and eastern United States, and (2) the old synonym 
Anatrytone Dyar, with eight species, ranging from southern Canada to Argentina. Half 
of these species were in Mellana (and in synonymy besides); and they include the type 
species of Mellana, which thus becomes a new synonym of Anatrytone. For much of 
what was in Mellana, plus five new species, I genitalically define the new genus Quasi- 
mellana, with a total of 24 species, ranging from the far southern United States to northern 
Argentina. Despite superficial similarities between some species, Anatrytone and Qua- 
simellana are not closely related. 

All names in Anatrytone, except those of the type species Anatrytone logan (Edwards) 
and the new species A. sarah, and all names in Quasimellana, except those of the new 
species Quasimellana siblinga, Q. antipazina, Q. andersoni, Q. imperfida, and Q. mielkei, 
are new combinations. The type species of Quasimellana is Q. mexicana (Bell). In the 
following lists, eight new synonyms appear in brackets. 

I genitalically define two species groups in Anatrytone, each of which (a) ranges widely 
(Canada to Argentina/Mexico to Argentina) and (b) comprises four species that tend to 
replace one another geographically: (1) the logan group with the North American logan 
subgroup comprising Anatrytone logan (Edwards) and A. mazai (Freeman) and the South 
American barbara subgroup comprising A. barbara (Williams & Bell) and A. flavens 
(Hayward); and (2) the compact mella group comprising A. potosiensis (Freeman), A. 
mella (Godman), A. sarah n. sp., and A. perfida (Moschler) [= gladolis (Dyar)]. 

I genitalically define three species groups in Quasimellana: (1) the eulogius group 
(southern United States to Brazil and Paraguay, but mainly North American) with the 
five species Q. mexicana (Bell), QO. eulogius (Plotz) [= agnesae (Bell), = oaxaca (Freeman)], 
Q. siblinga n. sp., Q. balsa (Bell) [= balsa freemani (Steinhauser)], and Q. mulleri (Bell); 
(2) the sethos group (Mexico to Bolivia and Brazil, and equally North and South American) 
with the 12 species Q. aurora (Bell) [= tecla (Steinhauser)], O. nayana (Bell) [= tamana 
(Steinhauser)], QO. noka (Evans), Q. pazina (Evans), Q. antipazina n. sp., Q. sista (Evans), 


274 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Q. andersoni n. sp., Q. sethos (Mabille), Q. myron (Godman), Q. verba (Evans), Q. 
inconspicua (Hayward), and Q. angra (Evans); and (8) the nicomedes group (Mexico to 
Argentina, but mainly South American) with the seven species Q. amicus (Bell), Q. fieldi 
(Bell) [= montezuma (Freeman)], Q. nicomedes (Mabille) [= monica (Pl6étz)], Q. imperfida 
n. sp., Q. mielkei n. sp., QO. meridiani (Hayward), and Q. pandora (Hayward). 

Four species put in Mellana by Evans belong neither in Quasimellana nor in Anatry- 
tone: gala Godman, helva Moschler, rivula Plotz, and clavus Erichson, which is the 
differentiate of Wallengrenia currently going by the junior name Wallengrenia otho 
curassavica (Snellen). 


Additional key words: systematics, species groups and sister species, New World, 
geographic distribution, Wallengrenia. 

Genitalia are the best means to an end—for the holder, of course, 
but also, at quite another level, for the thoughtful beholder. Long 
exploited for separating species from one another, genitalia are just as 
good for pulling related species together in higher groups like genera. 

Although Scudder and Burgess (1870, plus later work summarized 
in Scudder 1889) and Godman and Salvin (1879-1901) were way ahead 
of their time because they examined and illustrated the male. genitalia 
of many of the skipper butterflies that they treated, they erred by 
placing species with similar genitalia in different genera and species 
with disparate genitalia in the same genus. For example, the twelve 
species of Atrytone whose genitalia grace plate 94 in Godman and 
Salvin actually belong to five distinct genera. 

In North America, illustration of skipper genitalia continued with 
Barnes and McDunnough (1912), Skinner (1914), and especially Skinner 
and Williams (1922, 1923a, 1928b, 1924a, 1924b, 1924c) who figured 


the males of almost all the species then known from north of Mexico. 
When Lindsey, Bell, and Williams in 1931 updated Lindsey’s 1921 


treatment of this fauna, they incorporated the Skinner and Williams 
figures. | 

An overly guarded paragraph in Lindsey (1921:11) on the taxonomic 
uses of genitalia reappeared nearly verbatim in the collaborative re- 
vision (Lindsey et al. 1931:10), but with a couple of addenda (here set 
in italics): 


The genitalia, especially in the males, are of great value in making 
specific identifications and similarity of genitalic structure often 
affords an index of generic relationship. We have found several 
apparent contradictions of the latter statement and are therefore 
inclined to use the principle cautiously until more is known about 
the skippers, but still we hesitate to include in the same genus 
species whose genitalia are of widely different forms unless other 
structures indicate close relationship. Genitalia are obviously more 
erratic in their variation than other structures. 


These italic afterthoughts took a huge leap backward. 


VOLUME 48, NUMBER 4 275 


Though it soon became standard, in this country and abroad, to 
illustrate male genitalia in descriptions of new skipper species, the 
enormous store of information in published tails was—and is—relatively 
untapped. Authors would dutifully show a new tail but often say little 
or nothing about it, even with respect to the species from which it 
came, much less compare it critically with others for clues to higher 
affinities. Despite exploitation by Barnes and McDunnough (1912) in 
megathymines, female genitalia were mostly ignored until MacNeill 
(1964) in Hesperia and Burns (1964) in Erynnis made heavy, compar- 
ative use of them. Female genitalia can be as helpful as those of the 
male. 

We are still seduced by outward appearance: simius Edwards looks 
like an Amblyscirtes—but its genitalia, in both sexes, spurn the Am- 
blyscirtes mold; reexamination of ‘“‘other structures” shows that the palpi 
and especially the antennal apiculus also deviate significantly; and sim- 
ius must go elsewhere (Burns 1990). Again, nabokovi Bell & Comstock 
(1948) superficially looked like an Atalopedes to its describers, who 
routinely figured the tail of the male—but those genitalia, in both sexes, 
are unabashed variations on the Hesperia (not the Atalopedes) theme; 
the stigma, too, comes straight from Hesperia; and that is where na- 
bokovi goes (Burns 1987, 1989). Despite appearances (and convention), 
genitalia clearly show that snowi (Edwards) belongs in Paratrytone 
rather than Ochlodes and that two-thirds of the species currently in 
Paratrytone belong somewhere else—mostly with the non-marsh-dwell- 
ing species of Poanes, which, in turn, harbor a pair of outwardly ac- 
ceptable genitalic misfits (Burns 1992). 

Granted, genitalia are far more difficult, tedious, and time-consuming 
to study and compare rigorously. But their morphologic complexity in 
skippers yields a wealth of potential characters (often enriched through 
asymmetry). Being hidden and serving quite other functions, they es- 
cape many of the pressures bombarding the external, visual phenotype 
and tend to reflect relationship better than facies. 

During the last eight years I have been genitalically reviewing nearc- 
tic hesperiine skippers, a relatively well-studied fauna whose long-stable 
genera seem to be gaining authority—yea, rigidity—through constant 
repetition in a spate of state, provincial, seminational, and national 
butterfly books and checklists (e.g., Harris 1972, Irwin & Downey 1978, 
Hooper 1978, Shapiro 1974, MacNeill 1975, Dornfeld 1980, Stanford 
1981, Pyle 1981, Miller & Brown 1981, 1983, Opler & Krizek 1984, 
Scott 1986, Tilden & Smith 1986, Heitzman & Heitzman 1987, Shull 
1987, Klassen et al. 1989, Bailowitz & Brock 1991, Opler & Malikul 
1992, Iftner et al. 1992). Unfortunately, as I have intimated, our genera 
are a mess. Problems generally spread into the neotropics, where they 


276 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


ramify—but where specimens and data are so much scarcer that so- 
phisticated generic revision dealing fully with all relevant species is not 
practical. Even if it were, I am finding mistakes at the generic level 
too rapidly to give each affected species anything like the time and 
attention I have lavished on pairs of close and confusing differentiates 
within Erynnis, Celotes, Atrytonopsis, Autochion, Wallengrenia, and 
Pyrgus (Burns 1964, 1974, 1983, 1984, 1985, unpublished). Such detail 
is dispensable because problems of grouping species in genera are dif- 
ferent from problems of delimiting species and analyzing speciation. 

I aim here and in some related papers (e.g., Burns 1992, 1994) to 
redefine certain genera strictly by means of genitalia. Many short ge- 
neric descriptions—including those of Evans (1955)— that omit genitalic 
characters really fail to characterize. Some recent long descriptions 
manage to drown a few useful genitalic observations in an indiscrim- 
inate sea. At the very least, my corrected generic limits will lead to 
better evolutionary, ecologic, ethologic, and biogeographic generali- 
zations about American hesperiines. 

Since my examination of their genitalia has shown that some type 
specimens are not what others thought, I have had to make some jolting 
changes in nomenclature. The abbreviations I use in citing museums 
are spelled out at the beginning of Acknowledgments (p. 334). All 
X-rated genitalia dissections are mine. 


Atrytone Scudder vis-a-vis Anatrytone Dyar 
(Figs. 1-3, 16,17) (Figs. 4-15, 18-27, 83-86) 


Scudder (1872) “defined” his new genus Atrytone merely by des- 
ignating Hesperia iowa Scudder (= Hesperia arogos Boisduval & Le- 
conte) type species and by including three other species originally 
described in Hesperia: logan Edwards, conspicua Edwards, and za- 
bulon Boisduval & Leconte. Because this largely nearctic assemblage 
is heterogeneous, interpretation of Atrytone varied. Over the next eight 
decades, chiefly Godman, Dyar, Williams, Bell, and Hayward described 
many and various neotropical species in Atrytone. Hayward (1948) 
made one of them, mella Godman, the type of his monotypic new genus 
Mellana. Meanwhile, Barnes and McDunnough (1916) removed za- 
bulon (plus a few of its congeners) to Poanes; and, eventually, Evans 
(1955) removed conspicua (plus what by then were seven related con- 
geners) to Euphyes and all of the modern neotropical species of Atry- 
tone to Mellana, leaving Atrytone with just two nearctic species from 
the eastern and central United States and adjacent Canada: arogos and 
logan. Later, Freeman (1969) described two new species of Atrytone 
from Mexico: mazai and potosiensis. 

Dyar (1905) “characterized’’ his heterogeneous new genus Anatry- 


VOLUME 48, NUMBER 4 7 2H 


tone in four and a half lines of undiagnostic text, designated Hesperia 
delaware Edwards (= Hesperia logan) type species, and included also 
lagus Edwards (now considered a subspecies of logan), vitellius Fa- 
bricius (now in Choranthus), and, of all things, arogos—the type species 
of Atrytone! (Dyar placed in Atrytone four skippers congeneric with 
one another though not with arogos.) Skinner (1905) lost no time in 
pointing out this grand idiocy, and Anatrytone has been called a syn- 
onym of Atrytone ever since (e.g., Barnes & McDunnough 1916, Lind- 
sey 1921, Lindsey et al. 1981, Evans 1955, dos Passos 1964, Miller & 
Brown 1981, 1983). 

But the two genera do have different types, arogos and logan; and, 
as I demonstrated in great genitalic detail at the annual meetings of 
The Lepidopterists’ Society in 1990 and 1991, these two fairly similar 
looking species (which have always been grouped—even by Dyar) really 
belong in separate genera. We must refer to Atrytone arogos and 
Anatrytone logan. Seizing mostly on differences in immature stages 
and life history, Scott (1992) reached the same conclusion. 

For the record, I wish to emphasize that male tails of these skippers 
have been adequately shown in the past (though improperly read): 
logan by Scudder (1889:pl. 37, fig. 11) and, more completely, by God- 
man (1900:pl. 94, fig. 6); both arogos and logan by Skinner and Williams 
(1924a:figs. 12, 183—reprinted in Lindsey et al. 1931 on plates 27 and 
28). 

How do the genitalia of Atrytone and Anatrytone differ? Not with 
the “saccus longer” in Anatrytone, as claimed by Scott (1992:135). 

An answer requires more than comparing the type species. Although, 
with the loss of logan, Atrytone is currently monotypic, Anatrytone is 
decidedly polytypic, containing not just logan and the above mentioned 
Freeman species, mazai and potosiensis, but five other species that 
collectively range from Mexico to Paraguay and Argentina. Evans (1955) 
put four of those species in Mellana—and in synonymy besides! Scott 
(1992:186) wrongly asserted ‘that based on adult morphology ‘Atry- 
tone’ potosiensis Freem. obviously belongs to genus Mellana potosien- 
sis new combination.” In characterizing Anatrytone, I have studied 
and compared the genitalia of all eight species, and have chosen to 
figure fully those of potosiensis, barbara Williams & Bell, and flavens 
Hayward, as well as logan, because they are as different from logan 
as any in the genus. However, in showing extremes of genitalic variation 
within Anatrytone, I am more interested in conveying a sense of the 
resemblance among the species than of the differences between them. 
Once this essential similarity is grasped, the larger and qualitatively 
different gap between Anatrytone and Atrytone should be obvious. 

In males of Atrytone (Figs. 1-8), the valva is simple: its only em- 


278 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 1-3. Male genitalia of Atrytone arogos from loess hills, 1460 ft (445 m), 4% mi 
(7% km) SE Westfield, Plymouth County, Iowa, USA, 1 July 1980, J. M. Burns (genitalic 
dissection no. X-2569) (USNM). Scale = 1.0 mm. 1, Tegumen, uncus, and gnathos in 
dorsal view. 2, Complete genitalia (minus right valva), with vesica everted, in left lateral 
view. 3, Aedeagus, with vesica everted, in dorsal view. 


VOLUME 48, NUMBER 4 : 279 


Fics. 4-6. Male genitalia of Anatrytone logan from loess hills, 1150-1300 ft (850- 
395 m), 2.8 mi (4.5 km) W Ticonic, T85N, R44W, sect. 18 and 19, Monona County, 
Iowa, USA, 2 July 1980, J. M. Burns (X-2567) (USNM). Scale = 1.0 mm. 4, Tegumen, 
uncus, and gnathos in dorsal view. 5, Complete genitalia (minus right valva), with vesica 
everted, in left lateral view. 6, Aedeagus, with vesica everted, in dorsal view. 


bellishment is a small, toothlike flange on the distal dorsal rim (Fig. 2). 
The tegumen/uncus, in lateral view (Fig. 2), is dorsally concave above 
the point at which the gnathos diverges, and, in dorsal view (Fig. 1), 
is unevenly tapered to an extremely delicate pair of uncus prongs so 
that much of the underlying gnathos shows. The aedeagus (Figs. 2, 3) 
is short (distinctly shorter than the rest of the intact genitalia), stout, 
and distally flared. The everted vesica (Figs. 2, 3), which is short, sports 
two pairs of rigid cornuti below—the pair at the distal end of the 
aedeagus with more teeth than the pair beyond it. 

In males of Anatrytone (Figs. 4-15, 24-27), the valva is more com- 
plex: a large notch in the dorsal rim splits a prominent dorsodistal 


280 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


extension from the body of the valva (Figs. 5, 8, 11, 14, 27); toward 
and at the base of this extension are modest medial protrusions (Figs. 
5, 8, 11, 14). The tegumen/uncus, in lateral view (Figs. 5, 8, 11, 14, 
27), is dorsally convex above the point at which the gnathos diverges, 
and, in dorsal view (Figs. 4, 7, 10, 13), is more or less evenly tapered 
to a heavier pair of uncus prongs so that most (Fig. 4) or all (Figs. 7, 
10, 18) of the underlying gnathos is concealed. The aedeagus (Figs. 5, 
6, 8,9, 11, 12, 14, 15, 24-27) is nearly as long as, or longer than, the 
rest of the intact genitalia; and its floor is caudally prolonged. The 
everted vesica (Figs. 5, 6, 8, 9, 11, 12, 14, 15, 24-26), which is long, 
sports one pair of rigid cornuti below, near the caudally prolonged floor 
of the aedeagus, plus (in all species but barbara [Figs. 11, 12] and flavens 
[Figs. 14, 15]) a pair of flexible, spinulose cornuti above (which suggest 
scouring pads). 

The female genitalia look more immediately distinct: they are shorter 
in Atrytone than in Anatrytone. In females of Atrytone (Figs. 16, 17), 
the lamella postvaginalis is short (in ventral view, wider than long) and 
arched dorsad (see lateral view); its posterior margin is usually well 
notched at the midline. The ductus bursae is short, almost uniformly 
wide, and well sclerotized, with many longitudinal wrinkles, especially 
dorsally. The adjacent, posterior part of the corpus bursae is lightly 
sclerotized, with more (chiefly longitudinal) wrinkles. 

In females of Anatrytone (Figs. 18-21) other than barbara (Figs. 22, 
23) and perhaps flavens (whose female I have not seen), peculiar, 
roughly linear sclerotization occurs midventrally in membrane between 
the ovipositor lobes, posterior and dorsal to the lamella postvaginalis 
and separate from it. The lamella postvaginalis itself (Figs. 18-23) is 
long (in ventral view, longer than wide) and ventrally about flat, with 
its lateral edges bent dorsad (see both ventral and lateral views); its 
posterior margin varies from outwardly rounded to shallowly notched 
at the midline. The ductus bursae—though ranging from short to long 
and from well to poorly sclerotized—reflects a basic pattern that in- 
volves some tapering toward the ostium bursae (Figs. 18, 20, 22), some 
dorsal concavity (Figs. 19, 21, 23), and some tendency for its roof to 
become membranous anteriorly where it meets the corpus bursae. The 
corpus bursae is entirely membranous. 


THE SPECIES OF ANATRYTONE (INCLUDING THE TYPE OF MELLANA) 
The logan Group 
(Figs. 4-6, 10-15, 18, 19, 22, 23) 
Anatrytone logan is the northernmost member of a loose, four-species 
group extending from southern Canada to Peru, Brazil, and northern 
Argentina. | 


VOLUME 48, NUMBER 4 281 


In males of the logan group, the saccus is long (Figs. 5, 11, 14); 
titillators are totally absent (Figs. 12, 15) or, in about half the individuals 
of logan, present in rudimentary form on just the right side of the 
aedeagus (Fig. 6); the rigid cornuti are more or less linear and long and 
parallel with the caudal prolongation of the aedeagal floor so as to 
suggest dorsolateral extensions of the aedeagal roof (Figs. 5, 6, 11, 12, 
14, 15); and the valvae are usually low (Figs. 11, 14) to medium (Fig. 
5) in height. 

In logan group females (flavens not seen), the sclerotization in mem- 
brane between the ovipositor lobes is wide (Fig. 18) or, in barbara, 
absent (Figs. 22, 23); the lamella postvaginalis, moderately long; the 
ventral lip of the ostium bursae, relatively simple; and the ductus bursae, 
long (see Figs. 18, 19, 22, 23 for all the above characters). 

Although well differentiated from one another, these species are more 
or less allopatric: A. logan occurs from extreme southcentral Canada, 
through most of the eastern and central United States, to extreme 
northeastern Mexico; A. mazai, from extreme southern Texas (Laredo 
[Freeman 1969]) and northeastern Mexico, through El Salvador, to Costa 
Rica (Guanacaste); A. barbara, from Colombia, through Ecuador, Ven- 
ezuela, and the Guianas, to Peru and Brazil (Amapa, Para, Rondénia); 
and A. flavens, in northern Argentina (Tucuman, Salta). This far-flung 
quartet forms two subgroups of two species each—a North American 
logan subgroup comprising logan and mazai and a South American 
barbara subgroup comprising barbara and flavens. Anatrytone logan 
and A. mazai are closer to each other, both morphologically and geo- 
graphically, than are A. barbara and A. flavens. However, logan and 
mazai are not subspecies, as claimed by Scott (1986) and echoed by 
Opler and Malikul (1992). Indeed, though virtually allopatric, they are 
too different from each other even to be grouped in a superspecies. 

In A. logan and A. mazai, the dorsodistal extension of the valva, 
viewed from the side, is dorsally broad (Fig. 5); the flexible “scouring- 
pad” cornuti are large and long and well sclerotized (Figs. 5, 6); the 
straight rigid cornuti have at least two to as many as five points (Figs. 
5, 6); and the caudal prolongation of the aedeagal floor is blunt at its 
distal end (Fig. 6). 

This prolongation is greater in A. mazai than it is in A. logan; the 
rigid cornuti of mazai are more than twice as long as those of logan; 
and the thin, platelike protrusion from the inner side of the dorsodistal 
extension of the valva is much larger—and arises much higher—in 
mazai than in logan. 

In A. barbara and A. flavens, the dorsodistal extension of the valva, 
viewed from the side, is dorsally narrow (Figs. 11, 14); flexible cornuti 
are lacking (Figs. 11, 14); the long, curved rigid cornuti have either a 


282 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 7-9. Male genitalia of Anatrytone potosiensis from San Pedro Sula, HON- 
DURAS, 17 July 1979, R. D. Lehman (X-2573) (USNM). Scale = 1.0 mm. 7, Tegumen, 
uncus, and gnathos in dorsal view. 8, Complete genitalia (minus right valva), with vesica 
everted, in left lateral view. 9, Aedeagus, with vesica everted, in dorsal view. 


VOLUME 48, NUMBER 4 283 


Fics. 10-12. Male genitalia of Anatrytone barbara from 20 km SW Puerto Maldonado, 
300 m, Madre de Dios, PERU, 25 October 1983, S. S. Nicolay (X-2585) (USNM). Scale 
= 1.0 mm. 10, Tegumen and uncus in dorsal view. 11, Complete genitalia (minus right 
valva), with vesica everted and juxta stippled, in left lateral view. 12, Aedeagus (minus 
vesica) in dorsal view. 


single (terminal) point (Figs. 11, 12) or such a point plus one small 
accessory point (Figs. 14, 15); and the caudal prolongation of the ae- 
deagal floor is forked at its distal end (Figs. 12, 15). 

The saccus is far shorter in A. flavens (Fig. 14) than it is in A. barbara, 
where it is nearly or quite as long as the valva (Fig. 11); the gnathos is 


284 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


13 


15 


J_ 


Fics. 18-15. Male genitalia of Anatrytone flavens from Salta, ARGENTINA (X-3115) 
(Mielke collection). Scale = 1.0 mm. 13, Tegumen and uncus in dorsal view. 14, Complete 
genitalia (minus right valva), with vesica everted and juxta stippled, in left lateral view. 
15, Aedeagus (minus vesica) in dorsal view. 


much shorter than the uncus in A. barbara (Fig. 11) though not in A. 
flavens (Fig. 14); the dorsodistal extension of the valva, viewed from 
the side, is dorsally extra narrow in A. flavens (Fig. 14); the caudal 
prolongation of the aedeagal floor is shallowly forked, with four tines 
at its distal end, in A. flavens (Fig. 15) but deeply and widely forked, 


VOLUME 48, NUMBER 4 285 


with two main tines (often equipped with one [Fig. 12] or occasionally 
two small secondary teeth), in A. barbara (Fig. 12); the sides of the 
caudal prolongation are distinctively rolled upward in A. flavens (Fig. 
15) while the distal end of the aedeagus is more expanded in A. barbara 
(Fig. 12). 

In both A. logan and A. mazai, as already noted, there is wide 
sclerotization in membrane between the ovipositor lobes (Figs. 18, 19); 
the ventral lip of the ostium bursae is not notably enhanced (Figs. 18, 
19); the ductus bursae is not strongly flattened dorsoventrally, and its 
dorsal concavity is pronounced (Fig. 19); at most, the anterior quarter 
of the roof of the ductus bursae gives way to membranous corpus bursae 
(Fig. 19); and the tapering of the ductus bursae toward the ostium 
bursae looks prosaic (Fig. 18). 

The dorsal concavity of the ductus bursae is simple throughout its 
length in A. mazai whereas a more or less anterior part of the dorsal 
concavity is divided longitudinally by a middorsal keel in A. logan 
(Figs. 18, 19); the ventral lip of the ostium bursae is unreinforced in 
mazai but lightly reinforced in logan (Figs. 18, 19). 

In A. barbara, there is no sclerotization in membrane between the 
ovipositor lobes (Figs. 22, 23); the ventral lip of the ostium bursae is 
well reinforced and turned strongly downward and backward (Figs. 
22, 23); the ductus bursae is quite flattened dorsoventrally, and its dorsal 
concavity is slight (Fig. 23); at least the anterior half of the roof of the 
ductus bursae gives way to membranous corpus bursae (Figs. 22, 23); 
and the ductus bursae tapers toward the ostium bursae with style (Fig. 
ai The mella Group 

(Figs. 7-9, 20, 21, 24-27, 83-86) 


Anatrytone potosiensis is the northernmost member of a compact, 
four-species group extending from Mexico to Argentina. 

In males of the mella group, the saccus is short (Figs. 8, 27); con- 
spicuous, paired titillators project from the sides of the aedeagus about 
where the caudal prolongation of the aedeagal floor begins (Figs. 8, 9, 
24-27); the rigid cornuti are short (Figs. 8, 9, 24, 25) or lengthened by 
a peculiar base (Fig. 26); and the valvae are usually medium to extra 
high (Figs. 8, 27). 

In mella group females, the sclerotization in membrane between the 
ovipositor lobes is narrow; the lamella postvaginalis, extra long; the 
ventral lip of the ostium bursae, well arched caudally and heavily 
sclerotized; and the ductus bursae, short (see Figs. 20, 21 for all the 
above characters). 

Records of these closely related species indicate some sympatry with 
no breakdown in genitalic differences: Anatrytone potosiensis occurs 


286 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 16,17. Female genitalia of Atrytone arogos from Sioux City, Woodbury County, 
Iowa, USA, 26 June 1988, A. W. Lindsey (X-2745) (USNM). Scale = 1.0 mm. 16, Sterigma 
and bursa copulatrix in ventral view. 17, The same, plus part of the ductus seminalis, in 
right lateral view. 


from central Mexico, through Guatemala, to Honduras; A. mella, from 
southern Mexico, through Central America (Guatemala, Belize, Hon- 
duras, Costa Rica, Panama), to Colombia and Peru; A. sarah (described 
below) from southern Colombia, through Ecuador and Peru, to Bolivia; 
and A. perfida (Méschler), from Colombia, Venezuela, Trinidad, Guy- 
ana, and French Guiana, through Brazil, to eastern Peru, Paraguay, 
and northern Argentina. 

The following comparative description simultaneously characterizes 
all four species of the mella group; sarah is phylogenetically closest to 
mella. 


Anatrytone sarah, new species 
(Figs. 24, 83-86) 


Most readily told by the aedeagus, especially the paired titillators which are longer 
and distally recurved in sarah (Fig. 24) and mella (Fig. 25) but shorter and divergent in 


VOLUME 48, NUMBER 4 287 


Fics. 18, 19. Female genitalia of Anatrytone logan from Austin, Travis County, 
Texas, USA, 14 May 1966, J. M. Burns (X-2627) (USNM). Scale = 1.0 mm. 18, Sterigma 
and bursa copulatrix, plus linear sclerotization in membrane between ovipositor lobes, in 
ventral view. 19, The same, plus part of the ductus seminalis, in right lateral view. 


288 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fics. 20, 21. Female genitalia of Anatrytone potosiensis from Cotaxtla Exp. Sta., 
Cotaxtla, Veracruz, MEXICO, 31 July 1962 (X-2606) (USNM). Scale = 1.0 mm. 20, 
Sterigma and bursa copulatrix, plus linear sclerotization in membrane between ovipositor 
lobes and an indication of the terminal abdominal tergites (including the ovipositor lobes), 
in ventral view. 21, The same, plus part of the ductus seminalis, in right lateral view. 


VOLUME 48, NUMBER 4 289 


Fics. 22, 23. Female genitalia of Anatrytone barbara from Tambopata Reserve, Rio 
La Torre, 300 m, Madre de Dios, PERU, 2 November 1984, S. S. Nicolay (X-2829) 
(USNM). Scale = 1.0 mm. 22, Sterigma, bursa copulatrix, and an indication of the terminal 
abdominal tergites (including the ovipositor lobes) in ventral view. 23, The same, plus 
part of the ductus seminalis, in right lateral view. 


290 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


potosiensis (Figs. 8, 9) and perfida (Fig. 26). The titillators are shortest in perfida. In 
sarah (Fig. 24) they are abruptly constricted from a broad, knobby base and are bowed 
farther out than they are in mella (Fig. 25) whose basal taper is more even. They are 
also less flattened in sarah than in mella but lie more in a horizontal plane. In sarah, as 
in mella and most perfida, each titillator ends in a single point (Figs. 24-26), instead of 
in 2 to 4 (usually 3) points, as in potosiensis (Figs. 8, 9). (In one of the many males of 
perfida examined, the left titillator ends in 2 points.) 

The caudal prolongation of the aedeagal floor is in sarah long, generally somewhat 
narrower than in other members of the mella group, continuously tapered from front to 
back, and distally truncate (Fig. 24); it is also truncate in potosiensis but relatively short 
and broad (Fig. 9). Though long, it is distally rounded in mella (Fig. 25) and rounded 
(Fig. 26) to somewhat truncate in perfida—and usually centrally notched in perfida (Fig. 
26) but not in mella (Fig. 25). A lateral view shows it dipping downward sharply in sarah, 
less so in perfida, still less in mella, and little or not at all (Fig. 8) in potosiensis. Despite 
plenty of individual variation, these characters of the aedeagal floor are good. 

Development of both kinds of cornuti, taken together, is intermediate in sarah, greatest 
in perfida, least in mella and potosiensis (in different ways). The paired flexible cornuti 
(“scouring pads’) are large and well sclerotized in perfida (Fig. 26), middle-sized and 
well sclerotized in sarah and mella (Figs. 24, 25), and small and lightly sclerotized in 
potosiensis (Figs. 8, 9)—to the extent that in one individual they virtually vanish. The 
paired rigid cornuti are largest in perfida, where each has an extra-long base—more or 
less arcuate in dorsal (Fig. 26) or ventral view—and a long point (which becomes dual 
on the left side in one of the many perfida examined). The rigid cornuti are middle-sized 
in sarah, where each has a respectable base and one or sometimes two usually longish 
points (the two-point condition can affect the left or the right [Fig. 24] cornutus). The 
rigid cornuti are also middle-sized in potosiensis, where each has a base at least as large 
as that of sarah but usually a shorter point (Fig. 9). They are smallest in mella, where 
they either have smallish to (more often) small, narrow bases and longish to (more often) 
short points (Fig. 25), or else are drastically reduced to a sliver (such vestigial cornuti can 
be on the left, on the right, or on both sides). 

In ventral view the ventral lip of the ostium bursae forms a caudally directed arch, 
which is sharper in sarah and mella than in potosiensis (Fig. 20) and perfida. Although 
in all four species this lip curves downward (ventrad), it also curves conspicuously forward 
along both sides in sarah and mella but not in potosiensis (Figs. 20, 21) and perfida. 
Anterior to the lip region the ductus bursae is more or less lightly and incompletely 
sclerotized in mella, three-quarters to fully sclerotized in sarah, and fully sclerotized in 
potosiensis (Figs. 20, 21) and perfida. The arch of the ventral lip does not extend as far 
back in sarah, mella, and potosiensis as it does in perfida. 

Anatrytone sarah is similar in size to other species of Anatrytone (except barbara and 
especially flavens, which are small), with the usual sex difference (females larger than 
males): 9 66 average 15.5 mm (range 14.9-16.1 mm) in forewing length; 2 99, 16.5 mm 
(range 16.4-16.6 mm). 

The members of the mella group are similar (and, of course, individually variable) in 
superficial appearance—females so much that I cannot distinguish those of sarah (Figs. 
85, 86) from those of mella, perfida, and potosiensis (on an average, however, females 
of perfida have the dorsal light areas a little more extensive and the yellow of the broad, 
ventral light areas brighter and clearer, less darkened with rust). Males of sarah (Figs. 
83, 84) tend to be more distinctive because of the size and darkness of an irregular dark 
band in the dorsal forewing running obliquely from the base of the wing mainly through 
the upper part of the proximal half of space 1b, the proximal end of space 2, the length 
of the cell, and space 5, or spaces 4 and 5, to the wide dark border along the outer margin. 
This oblique dark band is variably, and usually less well, expressed in mella and especially 
perfida (some mella come close) and is completely missing from potosiensis. On the dorsal 
hindwing, males of sarah send a narrow, yellow orange ray along vein 1b to the outer 
margin (Fig. 83). Altogether, males of sarah approach the unusual, dorsal look of males 
of barbara of the logan group, except that other wing veins in sarah are still dark where 
they cross light areas (Fig. 83). 


VOLUME 48, NUMBER 4 291 


Fics. 24-26. Aedeagi, with vesicae everted, in dorsal view, belonging to three species 
of Anatrytone in the mella group (compare also Fig. 9). Scale = 1.0 mm. 24, Anatrytone 
sarah (holotype) from Limoncocha, 240 m, Napo, ECUADOR, 24-27 June 1980, C. V. 
Covell Jr. (H739) (USNM). 25, Anatrytone mella from Macaracas, Los Santos, PANAMA, 
22 December 1984, G. Small (X-2581) (USNM). 26, Anatrytone perfida from Sapucay, 
PARAGUAY, W. T. Foster (X-2575) (USNM). 


Holotype. 6; ECUADOR, Napo (province), Limoncocha, 240 m, 24-27 June 1980, C. 
V. Covell Jr.; S. S. Nicolay genitalia dissection H739; USNM. 

Paratypes. n = 8 6 2 2. BOLIVIA, Santa Cruz (department), Las Juntas, 250 m, J. 
Steinbach: November 1913, 1 6, J. M. Burns genitalia dissection X-2836, CMNH; December 
1913, 1 9, X-28483, CMNH. COLOMBIA, Amazonas (commissary), Leticia, 20 April 1946, 


292 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fic. 27. Godman’s figure of the male genitalia of Anatrytone mella; complete gen- 
italia (minus left valva and juxta) in left lateral view. Note that, with the left valva 
removed, Godman’s figure shows the inner surface of the right valva, whereas my figures 
show the outer surface of the left valva. 


L. Richter, 1 6, E. L. Bell genitalia dissection G1911, AMNH; Puerto Asis, Rio Putumayo, 
1 February 1969, S. S. Nicolay, 1 6, H484, USNM. ECUADOR, Napo (province): Archi- 
dona, 800 m, 13 October 1986, S. S. Nicolay, 1 6, H970, USNM; Coca, 350 m, 12 May 
1975, Lefebre, 1 6, X-3466, collection of O. H. H. Mielke; Pastaza (province), Puyo-Napo 
Road, km 25, 1100 m, 11 November 1988, D. H. Ahrenholz, 1 6, X-3204, collection of 
S. S. Nicolay. PERU, Loreto (department), 40 km NE Iquitos, 14-20 January 1991, J. 
Glassberg, 1 6, X-3175, USNM; Madre de Dios (department): Puerto Maldonado, 290 m, 
14 October 1988, S. S. Nicolay, 1 ¢, X-2607, USNM,; Boca Rio La Torre [=Tambopata 
Reserve], 300 m, 23 November 1983, G. Lamas, 1 4, X-2846, MUSM. 

Etymology. The specific name sarah, which hails to and from my wife, is a noun in 
apposition to the generic name Anatrytone. 


A Twist OF NOMENCLATURE AND A DASH OF WALLENGRENIA 
(Figs. 27-30) 


Besides the one of sarah, I have examined the types (also male) of 
perfida Moschler (1878) (ZMHB), gladolis Dyar (1914) (USNM), flavens 
Hayward (1940) (IML), potosiensis Freeman (1969) (AMNH), and ma- 
zai Freeman (1969) (AMNH). 

When Dyar (1914) described gladolis he designated ““Cotypes, one 
male, three females.’’ I have studied all four specimens, which are 
similarly labelled and conspecific, and have chosen the lone male as 
lectotype. It bears the following tags, most of which are printed: [1] 
Georgetown/Br. Guiana, [2] BredSpecimen, [3] HWBMoore/Coll, [4, 
handwritten] Larva on blades/of sugar cane, [5, red] Type No./18115/ 
U.S.N.M., [6, handwritten] Atrytone/gladolis/type Dyar, [7] GENI- 
TALIA NO./X-2831/J.M.Burns 1990. Although Dyar noted that gla- 
dolis is “nearest to A. mella,”’ his original description (confined to the 
appearance and spread of the wings) is too brief and superficial to 
distinguish gladolis from mella—or from a good many other hesper- 
iines, either. It turns out that gladolis Dyar (1914) is a (new) synonym 
of perfida Méschler (1878). | 

The original figure of the male genitalia of A. mella (Godman 1900: 


VOLUME 48, NUMBER 4 293 


30 


rr! 


Fics. 28-30. Male genitalia of holotype of Hesperia clavus Erichson from British 
Guiana (X-2933) (ZMHB), which is a species of Wallengrenia similar to, or the same as, 
W. otho. Scale = 1.0 mm. 28, Tegumen, uncus, and the very tip of the gnathos in dorsal 
view. 29, Complete genitalia (minus right valva) in left lateral view. 30, Aedeagus in 
dorsal view. 


pl. 94, fig. 16 [reprinted in this paper as Fig. 27]) shows the short saccus 
characteristic of the mella group; the long, recurved titillator, evenly 
tapered at the base (less obvious in this lateral view), characteristic of 
mella; the “scouring-pad”’ cornuti characteristic of all species of An- 
atrytone except barbara and flavens; and a valva with the characteristic 
Anatrytone shape, but extra high. Valval height increases out of pro- 
portion to valval length in the mella group, especially in mella, sarah, 
and potosiensis (Fig. 8), and somewhat less consistently in perfida. 
When Hayward (1948) described the new genus Mellana, he took 
as its type (and sole species) Atrytone mella Godman (1900), which he 
considered the same as Atrytone gladolis Dyar (1914). Since Hayward 
cited a specimen from northern Argentina (Misiones), his type species 
must really be perfida Méschler (1878) (= gladolis Dyar [1914]) instead 


294 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


of mella; but that makes no difference. Both are congeneric with An- 
atrytone logan (Edwards), and Anatrytone Dyar (1905) has priority 
over Mellana Hayward (1948). 

Evans (1955:354, 355) muddled his much expanded treatment of 
Hayward’s (1948, 1950b) monotypic Mellana by calling mella Godman 
(1900) a synonym of clavus Erichson (1848)—which thus, in Evans’s 
eyes, became the type of Mellana—and by listing gladolis Dyar (1914), 
barbara Williams & Bell (1931), and flavens Hayward (1940) as ad- 
ditional synonyms of clavus. I have shown that gladolis is a synonym 
of perfida, which is a species separate from mella but close to it, that 
barbara and flavens are very distinct from both of them and rather 
different from each other, and that all four go in Anatrytone. Having 
borrowed (from ZMHB) the holotype of Hesperia clavus Erichson 
(1848)—a male—and examined its genitalia (Figs. 28-30), I can flatly 
state that it is a species of Wallengrenia! 

Wallengrenia clavus (Erichson), new combination, is what Evans 
(1955 plus associated “Addenda and Corrigenda’’) called W. otho cur- 
assavica (Snellen). The name clavus, being older, supplants the name 
curassavica. The biologic entity clavus closely resembles otho Smith 
(1797), differing chiefly in the direction of the terminal tooth on the 
right (the longer) distal division of the aedeagus: that tooth points up 
and to the right in clavus, up and to the rear in otho (compare Figs. 
29, 30 with figs. 11-14, 19-22 in Burns 1985). The type of clavus comes 
from Guyana. In a crude transect—12 more tails of Wallengrenia males 
from Mexico to Brazil—the right aedeagal tooth points up and to the 
right in the six males from Panama, Colombia, Guyana, and French 
Guiana, but not in those from farther north or south. Some features of 
the pair of terminal, dependent, two-toothed cornuti (dubbed “‘flags”’ 
in Burns 1985) also appear to vary geographically. Whether clavus is 
a synonym of otho, a subspecies of otho, or a separate species is a 
problem beyond the scope of this paper. 

After blindly synonymizing barbara with three other species of An- 
atrytone and Wallengrenia clavus, Evans (1955:359) went on to de- 
scribe new species Mellana villa, which Mielke (1973) pegged as a 
synonym of M. barbara (Williams & Bell)! Mielke also observed that 
M. barbara was not synonymous with M. clavus (Erichson). 

Note that Atrytone, Anatrytone, and Quasimellana (the new genus, 
described below, for most of what was in Mellana) all lack a stigma 
on the forewing of the male whereas Wallengrenia bears a bold stigma. 


Summary of Atrytone and Anatrytone 


The following capsule of my treatment of Atrytone and Anatrytone 
gives the number of male and female genitalia examined at each level— 


VOLUME 48, NUMBER 4 : 295 


140 dissections in all. I list neither subspecies nor synonyms of Atrytone 
arogos and Anatrytone logan because I did not investigate these fa- 
miliar nearctic skippers at that low level. Such information is readily 
available in checklists (Miller & Brown 1981, 1983). Repeating it here 
would amount to empty endorsement of what may be faulty or un- 
documented opinion—the kind of deed that has left much of our tax- 
onomy a shambles. 


Atrytone Scudder, 1872, 10 6 15 2, n = 25 
arogos (Boisduval & Leconte, [1834]), 10 6 15 9 


Anatrytone Dyar, 1905, 82 6 33 2,n = 115 
= Mellana Hayward, 1948, new synonym 
the logan group, 31 614° 
the logan subgroup, 14 6 108 
logan (Edwards, 1863), 8 6 8 2 
mazai (Freeman, 1969), new combination, 6 6 2 ? 
the barbara subgroup, 17 ¢ 4 2 
barbara (Williams & Bell, 1931), new combination, 15 4 4 2 
= villa (Evans, 1955) 
flavens (Hayward, 1940), new combination, 2 ¢ 
the mella group, 51 6192 
mella (Godman, 1900), new combination, 14 6 11 ° 
sarah Burns, new species, 9 6 2 2 
potosiensis (Freeman, 1969), new combination, 6 6 1 2 
perfida (Moschler, 1878), new combination, 22 6 5 @ 
= gladolis (Dyar, 1914), new synonym 


Major Generic Characters in the Smallest Genitalic Structures 


What remains after subtracting species of Anatrytone (and Wallen- 
grenia clavus) from Mellana as treated by Evans (1955) is an unnamed 
and difficult neotropical genus at least thrice the size of Anatrytone 
and far more diverse. Ranging from the extreme southern United States 
(southern Texas; perhaps also southern Arizona—see Bailowitz & Brock 
1991) to Peru, Bolivia, Paraguay, and northern Argentina, new genus 
Quasimellana (described below) is broadly sympatric with Anatrytone. 
Both occur from sea level to moderate elevations. Despite some simi- 
larities in superficial appearance, which cause confusion, Quasimellana 
and Anatrytone are not closely related. 

We have seen that the genitalia of Anatrytone are basically conser- 
vative (in both sexes) and hence invaluable for defining that genus. In 
gross view, those of Quasimellana are signally variable, especially in 
males (females of a quarter of the species are not yet known): the large 
middorsal structures (tegumen/uncus plus underlying gnathos) and the 


296 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


MALE FOREWING LENGTH (mm) 


Ce a SSeS nr aN SS Std oa sg 
SESEELEZERS SELL SESS Eee as 
sgae2S5se2H 8 SRoOSSZEO0FaUsSeRSSSS 
Os E 5 wea & =) eal S| o— > oO 3 a - Sk9 —} 
”n = = 2 3 0 £ = oD Ben ae ok 
e°, ¢§s ZS x sao 3 Oo 
3 535 0 os 5 c= & Se 
SI S o 


QUASIMELLANA SPECIES 


Fic. 31. Size (male forewing length) in species of Quasimellana from smallest to 
largest. Vertical lines show ranges; dots, means (or single observations); crossbars, plus 
and minus twice the standard error of the mean (when the sample size exceeds five). 
Sample sizes appear above the range lines. There are two geographic subsamples of the 
widespread and relatively common Q. eulogius: eulogius-1 from Mexico and eulogius-2 
from Honduras, Nicaragua, Costa Rica, and Panama. 


large, paired lateral structures (valvae) assume too many forms to permit 
generalizing across the whole genus. But what do characterize Quasi- 
mellana are two relatively small and obscure genitalic structures cur- 
rently lacking the “respect and attention” (Burns 1987:184) they de- 
serve: the cornutus and the juxta. 

The shape of the cornutus alone is diagnostic. Though it does vary 
within—and more perceptibly between—species, the variation is so 


VOLUME 48, NUMBER 4 297 


Fics. 32-38. Cornuti of six species of Quasimellana showing variations on an essential 
theme. Scale = 0.5 mm. 32, Q. eulogius from Ciudad de Valles, San Luis Potosi, MEXICO, 
13 October 1976, E. C. Knudson (X-2587) (USNM). 33, Q. mexicana (X-2586) shown 
fully in Figs. 40, 41 (qv for specimen data). 34, Q. fieldi (X-2571) whose juxta appears 
in Fig. 39 (qv for specimen data). 35, Q. nicomedes from Cacatu, Antonina, 20 m, Parana, 
BRAZIL, 25 April 1973, Mielke (X-2366) (MacNeill collection). 36, Q. sethos from Paraiso, 
Canal Zone, PANAMA, 27 April 1982, S. S. Nicolay (X-2595) (USNM). 37, 38, QO. mulleri 
(paratype) from Guerrero, MEXICO, August, R. Miller (X-2669) (AMNH). 


298 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


TABLE 1. Number of antennal nudum segments in species of Quasimellana. 


Number of nudum segments 
Species groups EEE EEE —E— Eee 
and species Mean ll 12 13 14 15 16 n 


eulogius group 


mexicana 13.3 19 8 27 
eulogius 13.4 1 6 56 89 5 1 108 
siblinga 14.3 1 9 6 16 
balsa 13.4 1 9 6 1 17 
mulleri 12.9 Il 9 10 
sethos group 
aurora NB. 7 9 12 2 23 
nayana 14.4 1 3 4 8 
pazina 2 2 
antipazina 1 1 
sista ] ] 
andersoni 14.8 i 4 5 
sethos 14.0 4 16 I IL 22 
myron 12.9 i 10 11 
verba 13.5 4 4 8 
inconspicua 13.4 I 3 4 8 
angra 14.2 2 4 4 10 
nicomedes group 
amicus 13.2 8 2 10 
fieldi 14.2 1 3 42 17 1 64 
nicomedes eye 1 3 2 6 
imperfida 14.8 I 4 5 
mielkei 2 2 
meridiani WS 2) 2 4 
pandora 5) 5) 6 5 11 


constrained that a single, simple generic theme stands out (Figs. 32- 
38). Since the sclerotized cornutus sits in the membranous vesica which, 
except during copulation, rests collapsed and folded within the scler- 
otized aedeagus, the vesica must be everted for a really good look at 
the cornutus (Figs. 41, 48, 45, 47, 49, 50, 52, 53, 55, 57, 59, 60, 62, 65, 
67). 

The shape of the juxta (Fig. 39) is likewise diagnostic; but so, too, is 
its location. An apparent support for the aedeagus, the juxta lies ventral 
and lateral to it at about the level of the anterior ends of the paired 
valvae and the adjacent vinculum to which they articulate. Typically, 
in a lateral view of male genitalia, the juxta is largely to entirely hidden 
behind the vinculum and the front end of a valva. That is where it is 
in Atrytone (Fig. 2), Anatrytone (Figs. 5, 8, 11, 14), and Wallengrenia 
(Fig. 29); in many other genera thought to be related to these (i.e., in 
the Hesperia subgroup of the M or Hesperia group of Evans 1955), 
such as Hesperia itself (Burns 1987, fig. 2), Atalopedes (Burns 1989, 


VOLUME 48, NUMBER 4 299 


oy, 


Fic. 39. Juxta, in left anterodorsolateral view, of Quasimellana fieldi from 15.4 mi 
(24.8 km) S Ciudad de Valles, San Luis Potosi, MEXICO, 22 August 1967, G. F. Hevel 
(X-2571) (USNM). 


figs. 25, 27, 29, 31, 33), Polites (Burns 1994, fig. 14), Poanes (Burns 
1992, figs. 2, 5), Paratrytone (Burns 1992, fig. 20), and the like; and in 
a great many other skippers, as well. However, in Quasimellana the 
juxta is so far forward that most of it shows in lateral view (Figs. 41, 
48, 45, 47, 52, 55, 57, 59, 62, 65, 67). 

Since the location of the juxta is crucial, I regret to say that it can 
rarely be determined from published figures of skipper genitalia. In 
most of them, the juxta does not even appear (see Fig. 27, which 
exemplifies the many genitalic figures in Godman & Salvin 1879-1901). 
When it does, it almost always accompanies the aedeagus in an exploded 
view of the genitalia, displaced from its true position and from its 
proper relation to other parts. The common practice of dissociating 
major structures such as the valvae or the aedeagus when dissecting 
male genitalia, often destroys the juxta. Evans’s mode of quick dissection 
and dissociation routinely did away with it (Figs. 49, 50, 58, 60). 


Quasimellana, new genus 


(Figs. 31-82, 87-112) 


Size. Male forewing length ranges from 11.4 to 17.9 mm; but, in most species, mean 
male wing length falls between 14 and 16 mm (Fig. 31). The relatively few females 
available always average larger than conspecific males. 

Nudum. Ranging from 11 to 16 segments, the nudum usually numbers 13 to 15 segments 
(though often to 16 segments in the nicomedes group), evenly split between the body of 
the club and the apiculus (Table 1). 

Stigma. Males do not have a stigma. 

Male genitalia. The sole (or only sizable) rigid cornutus consists of a simple body with 
a head-and-neck that ends in a tiny hook (Figs. 832-38, 41, 48, 45, 47, 49, 50, 52, 53, 55, 
57, 59, 60, 62, 65, 67). The entire structure is well sclerotized and conspicuous. 

The juxta resembles a U whose paired, lateral, upright arms twist once into more or 


300 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


less expanded and posteriorly trending upper ends while the base of the U gives off a 
long, midventral, posteriorly tapering triangle (Fig. 39). The juxta sits unusually far 
forward, largely exposed in lateral view (Figs. 41, 48, 45, 47, 52, 55, 57, 59, 62, 65, 67). 
Female genitalia. The posterior margin of the lamella postvaginalis gives rise mid- 
ventrally to a single, posteriorly directed, bristly projection (Figs. 68-82). This central 
projection may be only slightly developed (Fig. 76) or remarkably wide (Fig. 74), but 
usually it is narrow and long (Figs. 68-73, 75, 77, 81, 82) to very long (Figs. 78-80). 
Type species. Atrytone mexicana Bell (1942b:461, fig. 5). Figs. 33, 40, 41 (male 
genitalia); 68, 69 (female genitalia); 87, 88 (adult male); and 89, 90 (adult female). 


Individually rather plain, the male genitalia are so interspecifically 
variable that they seem elaborate in Quasimellana as a whole. The 
tegumen/uncus varies from narrow to wide. The distal end of the uncus 
varies from about as thick as, to much thicker than, the distal end of 
the gnathos; from decidedly longer to decidedly shorter than the gna- 
thos; from close above the gnathos to far above it; from undivided to 
well divided; the uncal divisions, from closely parallel to widely di- 
vergent—and then, in one case, bent sharply upward, besides. The 
distal end of the valva may have zero, one, or two projections, each 
ranging from broadly rounded to sharply pointed and extending pri- 
marily dorsad or caudad (sometimes mediad, as well). The inner side 
of the valva may develop a triangular middle process, a more oblong 
dorsal process, both of these, or neither. The aedeagus is a simple tube 
(devoid of titillators and terminal teeth or prongs) about as long as the 
total intact genitalia (i.e., saccus plus valvae), but its vesica varies from 
long, narrow, and fingerlike to shorter and less regular, with protruding, 
armlike pouches; the diagnostic cornutus, situated near or at the distal 
end of the everted vesica, may or may not be accompanied by a tiny, 
secondary, apparently vestigial cornutus near the proximal end of the 
vesica. The diagnostic cornutus itself varies from short to long and from 
narrow to wide; in the shape, height, point of attachment, and angle 
of the head-and-neck on the body; and the body varies from convex 
to concave, as well as in certain minor decorations. The saccus is usually 
much shorter than the valva but may approach it or even exceed it in 
length. The juxta varies in just how far forward it sits, in the height of 
the twist in each arm, and in details of shape. 

Each species account begins (after any synonymy) with two or three 
kinds of crucial information in condensed form: (1) mention of the 
holotype, its sex, and its location (abbreviated, for example, “HTé- 
USNM”’?) whenever I have seen and studied that specimen and its 
genitalia; (2) the spatial distribution (based on material examined), 
arranged geographically by country (and, in the United States, Mexico, 
Brazil, and Argentina also by state, territory, or province), plus a few 
important additions from the literature; and (8) the numbers of males 
and females examined, followed (in parentheses) by the number of 
genitalia compared in each sex. Whenever I have studied the holotype 


VOLUME 48, NUMBER 4 301 


Fics. 40, 41. Male genitalia of Quasimellana mexicana from San Luis Potosi, San 
Luis Potosi, MEXICO, 3 August 1941, A. H. Moeck (X-2586) (USNM). Scale = 1.0 mm. 
40, Tegumen, uncus, and gnathos in dorsal view. 41, Complete genitalia (minus right 
valva), with vesica everted, in left lateral view. 


of a synonym, hyphenated abbreviations as in (1) end the entry in the 
synonymy. 

I have examined the genitalia of the types of two species (besides 
Wallengrenia clavus) put in Mellana by Evans (1955) but belonging 
neither in Quasimellana nor in Anatrytone: gala Godman (1900) 
(USNM), from Mexico, which Evans questioningly listed as a synonym 
of monica Plotz (1886), from southern Brazil (Santa Catarina); and 
rivula Mabille (1891) (ZMHB), which Evans treated as a species with 
two subspecies—rivula, from northern Brazil (Amazonas), and amicus 
Bell (1942), from Ecuador (although rivula is not a species of Quasi- 
mellana, amicus is). 

Though I amassed some 450 specimens in my study of Quasimellana, 
only 12 out of 24 species are represented by more than 10 specimens, 
but 7 of those are represented by more than 20; another 7 species are 
known from 6 or fewer specimens, and 8 of these (including 2 that 
Evans described back in 1955), from just 1. In light of these figures, 
the large (essentially neotropical) range of the genus, and the superficial 
similarities among various differentiates, additional species doubtless 


302 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fics. 42, 43. Male genitalia of paratype of Quasimellana siblinga from E] Vado-San 
Sebastian, 5500-6500 ft (1675-1980 m), ca 16° 53’ N, 96° 53’ W, Oaxaca, MEXICO, 22 
June 1992, J. Kemner (X-3592) (USNM). Scale = 1.0 mm. 42, Tegumen, uncus, and 
gnathos in dorsal view. 43, Complete genitalia (minus right valva), with vesica everted 
and juxta and cornuti stippled, in left lateral view (plus primary cornutus in profile). 


await discovery; and for most described species, knowledge of geo- 
graphic distribution leaves much to be desired. 


THE SPECIES GROUPS AND SPECIES OF QUASIMELLANA 


The eulogius Group 
(Figs. 31-33, 37, 38, 40-45, 68-78, 87-92) 


Male genitalia. Juxta: The juxta is posterior in location (Figs. 41, 43, 45), as it is in 
nayana, aurora, and andersoni of the sethos group (Figs. 47, 55). The level of the twist 
in each lateral arm of the juxta is low (Figs. 41, 43, 45), as it is in the sethos group (Figs. 
47, 52, 55, 57, 59). Cornutus: Except in mexicana (Figs. 33, 41), the head-and-neck of 
the cornutus is short, wide, and upright (Fig. 37) to backswept (Figs. 32, 43, 45). Its 
position on the body of the cornutus varies from central (Figs. 33, 41) to left of center 
(Fig. 38) to left (Figs. 32, 43, 45). The body of the cornutus is narrow except in mulleri 
(Figs. 37, 38). The tiny, secondary, vestigial cornutus is present (Figs. 41, 48, 45), except 
in two examined individuals of mulleri and one of siblinga, as it usually is in the sethos 
group (Figs. 47, 49, 52, 55, 57, 60). Vesica: The everted vesica is relatively short (Figs. 
41, 48, 45). Valva: On its inner surface, the valva always has a middle process, which is 
relatively large and long (Figs. 41, 43, 45), but never the dorsal process that marks the 
sethos group (Figs. 49, 50, 52, 53, 55, 57, 59, 60). In lateral view the dorsodistal end of 
the valva is simple and more or less broadly rounded (Figs. 41, 43, 45), without prominent 
sharp points or projections. Uncus: The distal end of the uncus is deeply divided, with 
the resultant prongs more or less close together and parallel (Figs. 40, 42, 44). In lateral 
view, the distal end of the uncus is only a little thicker than the distal end of the gnathos 


VOLUME 48, NUMBER 4 3 303 


44 


Fics. 44, 45. Male genitalia of Quasimellana balsa from 28 mi (87 km) S Ixtapan de 
la Sal, Guerrero, MEXICO, 16 August 1981, J. A. Chemsak (X-2803) (UCB). Scale = 1.0 
mm. 44, Tegumen, uncus, and gnathos in dorsal view. 45, Complete genitalia (minus 
right valva), with vesica everted and juxta and cornuti stippled, in left lateral view. 


(Figs. 41, 48, 45), as it is in the sethos group (Figs. 47, 49, 52, 55, 57, 59) plus amicus of 
the nicomedes group. 

Female genitalia. The anterior apophyses are long (Figs. 68, 69, 72, 73) except in balsa 
(Figs. 70, 71), where they are intermediate to short (the female of siblinga is unknown). 
The corpus bursae is relatively long and narrow (Figs. 68-73). 

General features. Sexual dimorphism is strong (Figs. 87-90), except in mulleri. Males 
are always extensively yellowish-orange and blackish-brown dorsally, with dark veins. 
The species are medium to large in size (Fig. 31). 

Distribution. Although the eulogius group (with 5 species) extends from the United 
States (south Texas) to Brazil and Paraguay, it is mainly North American. 


Quasimellana mexicana (Bell, 1942b:461, fig. 5), 
new combination 
(Figs. 31, 33, 40, 41, 68, 69, 87-90) 


HT6-USNM. Mexico (San Luis Potosi, Veracruz, Nayarit, Guerrero, Oaxaca), Guate- 
mala, Belize. 29 6 8 2 (16 6 4 9). 

The largest species of Quasimellana (Fig. 31), mexicana has a narrow cornutus with 
a distinctive, long, narrow head-and-neck obliquely stretched far in front of the body 
(Figs. 33, 41). The saccus is relatively long for Quasimellana (even for mexicana, however, 
the saccus of the individual in Fig. 41 is unusually long). The sclerotized lateral lobes of 
the lamella postvaginalis (on either side of the midventral, posteriorly directed, bristly 
projection) do not extend as far caudad as they do in most species of Quasimellana, while 
the ductus bursae is unusually plain (compare Figs. 68, 69 with Figs. 70-82). 


304 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Quasimellana eulogius (Pl6tz, 1883:64), new combination 
(Figs. 31, 32, 72) 


mellona (Godman, 1900:493, pl. 94, figs. 17-19). 

heberia (Dyar, 1914:5), lectotype 6 (J. M. Burns genitalia dissection X-2832) here 
designated from 3 cotypes (2 6 1 2 bred from larvae on blades of sugarcane, George- 
town, British Guiana, H. W. B. Moore, type no. 18116) in USNM. 

agnesae (Bell, 1959:18, figs. 8, 21), new synonym, HTé-AMNH. 

= oaxaca (Freeman, 1979:10, figs. 20, 21, 29 [valva upside down]), new synonym, HTé- 

AME. 


United States (south Texas—Cameron and Hidalgo counties), Mexico (Nuevo Leon, 
San Luis Potosi, Sinaloa, Veracruz, Puebla, Guerrero, Oaxaca, Chiapas, Yucatan, Quintana 
Roo), Guatemala, Honduras, Nicaragua, Costa Rica, Panama, Colombia, Venezuela, Trin- 
idad, Guyana, Brazil (Para, Espirito Santo); Evans (1955) also records Bolivia and Para- 
guay. 80 4 30 2 (49 4 29 9). 

For Quasimellana, eulogius is unusually common and widespread—the seeming “weed 
species’ of the genus. The large size and latitudinal and altitudinal diversity of my eulogius 
sample may explain why it showed the most individual variation (reflected, for example, 
in Fig. 31 and Table 1). But then again, it may include more than one species; and some 
of the above synonyms—particularly heberia and agnesae—may need resurrection. 

Godman (1907) promptly sank his mellona Godman (1900) to eulogius Plétz (1883). 
In describing agnesae (from two males taken 6 and 9 December 1939 in Acapulco, 
Guerrero, Mexico), Bell (1959:13) called it “a small species of Mellana, with different 
genitalia from those of its nearest relatives.” I have examined the holotype and paratype 
and Bell’s slides of their genitalia, which look just like those of eulogius. Although the 
small size (forewing length 12.2 and 12.7 mm) and greater extent of yellow orange coloring 
do differ from ordinary eulogius (Fig. 31), I have measured an equally dwarf (12.6 mm) 
but normal looking eulogius male from Guatemala and have seen normal sized but equally 
yellow orange eulogius males from Sinaloa, Mexico, and Brownsville, Texas. At least for 
now, I conservatively view the specimens of agnesae as light runts of eulogius. 

See siblinga for characterization of the male genitalia of eulogius (which are well shown 
in lateral view—including the extra long, narrow cornutus [Fig. 32]}—in Godman 1900: 
pl. 94, fig. 19). | 

The female genitalia (Fig. 72) depart from those of other known Quasimellana females 
because the ductus bursae is extremely short, and its main sclerotization (in ventral view) 
is a narrow band around it; the midventral, posteriorly directed, bristly projection from 
the posterior margin of the lamella postvaginalis is usually fairly long and narrow, without 
any laterally expanding sclerotization at its base; and the sclerotized lateral lobes that 
flank this projection are relatively narrow. (The female genitalia assigned to a paratype 
of Mellana tecla Steinhauser and to undetermined females of Mellana in figs. 82, 88, 
and 89 of Steinhauser 1974 all come from eulogius; even a delicate, female character 
state that I broadly ascribe to the eulogius group—long anterior apophyses—shows in 
these photographs. ) 

At present, I can cleanly pull the following species from a possible eulogius complex. 


Quasimellana siblinga, new species 


(Figs. 31, 42, 43, 91, 92) 


HT8-USNM. Mexico (Sonora, Sinaloa, Oaxaca). 16 ¢ (9 4). 

Superficially and genitalically reminiscent of eulogius and balsa. Primary cornutus 
(Fig. 43) similarly linear and narrow, about as long as in balsa (Fig. 45)—therefore much 
shorter than in eulogius (Fig. 32)—but body of cornutus dorsally convex and without the 
longitudinal twist of both eulogius (Fig. 32) and balsa (Fig. 45). Secondary, vestigial 
cornutus (Fig. 43) smaller than in eulogius (completely gone in one individual). In lateral 
view, dorsodistal end of valva somewhat truncate dorsally (Fig. 43) and so less rounded 
than in eulogius and balsa (Fig. 45). Ventrodistal corner of valva (Fig. 43) not cut away 


VOLUME 48, NUMBER 4 305 


as in eulogius. Tegumen, uncus, and gnathos (as well as distal end of aedeagus) (Figs. 
42, 43) much less massive than in balsa (Figs. 44, 45), and uncus prongs (Fig. 42) less 
widely notched than in balsa (Fig. 44)—all about as in eulogius; but each uncus prong 
proximally with a ventrolateral swelling (Figs. 42, 43)—not present in eulogius—large 
enough to show in dorsal view (Fig. 42). Gnathos slightly longer than uncus (Figs. 42, 
43). Medium sized species, close to fellow groupies other than mexicana (Fig. 31): mean 
male forewing length and SE = 14.81 + 0.14 mm, range 14.0-15.6 mm, n = 15. Usually 
14 or 15 nudum segments; on average (14.3), about one segment more than in fellow 
groupies (Table 1). 

Holotype. 6; MEXICO, OAXACA, El Vado-San Sebastian, ca 16° 53’ N, 96° 53’ W, 
5500-6500 ft (1675-1980 m), 22 June 1992, J. Kemner; USNM. 

Paratypes. n = 15 4, all from MEXICO. OAXACA (all collected by J. Kemner and 
housed in USNM): same data, 6 6, J. M. Burns genitalia dissections X-3590, X-3591, X- 
3592; same data except 27 June 1992, 2 6; 3-13 mi (5-21 km) N Sola de Vega and road 
to Grutas de San Sebastian, 5500-6500 ft (1675-1980 m), 6 July 1991, 1 6; same data 
except 6500 ft (1980 m), 1 4, J. M. Burns genitalia dissection X-3241; road to Grutas de 
San Sebastian, 6500 ft (1980 m), 10 July 1991, 2 6, H. A. Freeman genitalia dissections 
H-1197, H-1202; road to Grutas de San Sebastian, ca 16° 37’ N, 96° 57’ W, 5500-6500 ft 
(1675-1980 m), 10 July 1991, 1 6, J. Kemner & Romack, J. M. Burns genitalia dissection 
X-3589. SINALOA: Loberas Summit, 5 mi (8 km) NE Potrerillos, 1820 m, parkland forest, 
19 August 1973; 1 6, L. D. & J. Y. Miller, S. R. Steinhauser genitalia dissection SRS-2691, 
AME. SONORA: 13 mi (21 km) E El Novillo, 11 August 1985, 1 6, J. P. Brock, S. R. 
Steinhauser genitalia dissection SRS-1851, AME. 

Etymology. Quasimellana siblinga is a sibling species. 


Quasimellana balsa (Bell, 1942b:460, fig. 4), new combination 
(Figs. 31, 44, 45, 70, 71) 


= balsa freemani (Steinhauser, 1974:20, figs. 53-56, 85, 86), new synonym, HTé-AME. 


HTé-AMNH. Mexico (Sonora, Jalisco, Colima, Michoacan, Guerrero); Steinhauser (1974, 
1975) reports this species from El Salvador. 17 6 4 2 (14 6 4 2). 

See siblinga for definition of the male. 

Much as in mexicana (Figs. 68, 69), the sclerotized lateral lobes of the lamella post- 
vaginalis (on either side of the midventral, posteriorly directed, bristly projection) do not 
extend as far caudad (Figs. 70, 71) as they do in most species of Quasimellana; but the 
ductus bursae is remarkably short and oval (in ventral view [Fig. 70]), with an ostium 
(also somewhat oval) that opens far back at the base of the midventral, bristly projection, 
which is shorter and wider than it is in mexicana (see also Steinhauser 1974:fig. 86). 

The subspecies freemani is not helpful. Mellana balsa freemani was described from 
six males and three females all from one locality (Santa Tecla) in El Salvador (Steinhauser 
1974). When this short series was compared with the still shorter type series of balsa, 
amounting to four males from Guerrero, Mexico, it was said that “freemani males differ 
from typical balsa mainly in size and color” (Steinhauser 1974:22), the male genitalia 
being “essentially identical.” In every large sample of a Quasimellana species (and 
sometimes even in small ones) I have seen too much variation in color and pattern to 
give weight to the two color differences offered. Though real and noteworthy, the larger 
size of E] Salvador specimens does not warrant a formal name. By my measurements the 
type series of balsa is small, averaging 14.3 mm in forewing length (ranging from 13.7 
to 14.6 mm); but my total Mexico sample of balsa (n = 16), which includes those small 
types, is nearly a millimeter larger, averaging 15.04 mm (ranging from 13.7 to 15.7 mm) 
(Fig. 31). The El Salvador sample picks up where the Mexico sample leaves off and adds 
about another millimeter to the mean: according to Steinhauser (1974:22), the six freemani 
males average 16.2 mm (ranging from 15.8 to 16.5 mm). Elsewhere in Quasimellana, 
with enough eulogius to take good geographic subsamples, the forewings of 32 males 
from Honduras, Nicaragua, Costa Rica, and Panama averaged about half a millimeter 
more than those of 32 males from Mexico (Fig. 31). In Autochton cellus, wing length 
rose strikingly from central Mexico to southern Mexico to Guatemala and El Salvador 


306 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


(Burns 1984:10 and table 1). Skipper populations of a single species often increase in 
average wing length from north to south in the northern hemisphere (see tables and text 
in Burns 1964, 1984 for examples from Erynnis and Wallengrenia). 


Quasimellana mulleri (Bell, 1942b:462, fig. 6), 
new combination 
(Figs. 31, 37, 38, 73) 


HT8-USNM. Mexico (Tamaulipas, Durango, Jalisco, Colima, Guerrero). 935 2(768 
Q). 

The body of the cornutus is wide, with numerous peripheral wrinkles (Figs. 37, 38); 
and the rounded dorsodistal end of the valva tends to develop a modest point antero- 
dorsally. Overall the female genitalia (Fig. 73) look most like those of mexicana (Figs. 
68, 69); but the lateral lobes of the lamella postvaginalis extend well back as they do in 
most species of Quasimellana, while the midventral, posteriorly directed, bristly projection 
is about as delicate as it is in andersoni (Fig. 75), myron (Fig. 77), verba, inconspicua, 
and angra of the sethos group. 

Freeman (1967) argued that mulleri, which Evans (1955) questioningly called a syn- 
onym of eulogius, is really a distinct species. 


The sethos Group 
(Figs. 31, 36, 46-60, 74-77, 93-108) 


Male genitalia. Juxta: The juxta may be posterior (Figs. 47, 55) or anterior (Figs. 52, 
57, 59) in location. The level of the twist in each lateral arm of the juxta is low (Figs. 47, 
52, 55, 57, 59), as it is in the eulogius group (Figs. 41, 43, 45). Cornutus: The head-and- 
neck of the cornutus is relatively long and/or narrow (Figs. 36, 47, 49, 50, 52, 55, 57, 59, 
60), it projects obliquely forward from the body of the cornutus (Figs. 36, 47, 49, 50, 52, 
55, 59, 60), and its “dewlap’”’ is finely crenulate (Figs. 47, 49, 50, 52, 55, 59, 60). (Among 
other species of Quasimellana, only amicus has a crenulate “dewlap.’’) The body of the 
cornutus is dorsally flattish to concave (at some angles, concavity gives a “swaybacked”’ 
look—Figs. 36, 49, 52, 55, 59). The tiny, secondary, vestigial cornutus is present (Figs. 
47, 49, 52, 55, 57, 60), as it usually is in the eulogius group, except in the two known 
individuals of pazina (Fig. 50), the one of sista (Fig. 53), one of the examined males of 
verba, nearly half the examined males of inconspicua (Fig. 59), and most examined males 
of myron. Vesica: The everted vesica is relatively long (Figs. 47, 49, 50, 52, 58, 55, 57, 
59, 60). Valva: On its inner surface, the valva has at least a rudimentary, but usually a 
well developed, dorsal process, which is unique among species of Quasimellana (Figs. 
49, 50, 52, 58, 55, 57, 59, 60 [in these last two species, the dorsal process is short and 
extends mostly mediad, so it hardly shows in lateral view]). (The only males of the sethos 
group really lacking a dorsal process are two of the eight nayana examined, one of which 
appears in Fig. 47.) Although the distal end of the valva varies greatly, its basic pattern 
apparently is a pair of projections, one dorsal and the other more or less midcaudal. 
Uncus: The distal end of the uncus may be very well divided (Figs. 46, 48, 51) or undivided 
(Figs. 54, 56, 58). (Among other species of Quasimellana, only amicus has an undivided 
uncus.) In lateral view, the distal end of the uncus is only a little thicker than the distal 
end of the gnathos (Figs. 47, 49, 52, 55, 57, 59), as it is in the eulogius group (Figs. 41, 
43, 45) and in amicus of the nicomedes group. 

Female genitalia. The anterior apophyses are short (Figs. 74, 75) except in sethos (Fig. 
76), where they are intermediate (females of several species are unknown). The corpus 
bursae is relatively long and narrow (Figs. 74-77). 

General features. Sexual dimorphism is weak to strong. The species are small to large 
in size (Fig. 31). 

Distribution. Extending from Mexico to Bolivia and Brazil, the sethos group (with 12 
species) is equally North and South American. 


VOLUME 48, NUMBER 4 307 


Fics. 46, 47. Male genitalia of Quasimellana nayana from Santo Domingo, 15 mi 
(24 km) SE Simojovel, Chiapas, MEXICO, 8-15 July 1958, J. A. Chemsak (X-2789) 
(USNM). Scale = 1.0 mm. 46, Tegumen and uncus in dorsal view. 47, Complete genitalia 
(minus right valva), with vesica everted and juxta and cornuti stippled, in left lateral 
view. 


The nayana Subgroup within the sethos Group 
(Figs. 31, 46, 47, 74) 


Male genitalia. Juxta: The juxta is posterior in location (Fig. 47). Cornutus: The head- 
and-neck arises from the right side of the body of the cornutus. Uncus: The distal end 
of the uncus is very well divided, and the resultant uncus prongs are widely separated 
from each other (with a U-shaped gap) and somewhat divergent (Fig. 46). Gnathos: In 
lateral view, the gnathos is uniquely far below the uncus and uniquely short relative to 
the uncus (Fig. 47). 

Female genitalia. The midventral, posteriorly directed, bristly projection from the 
posterior margin of the lamella postvaginalis is hypertrophied and the rounded lobes of 
the lamella postvaginalis that flank it are atrophied (Fig. 74) so that the usual size 
relationships between these parts (Figs. 68-73, 75-82) are reversed. 

General features. Sexual dimorphism is strong. The species are medium sized (Fig. 
31). 

Distribution. The nayana subgroup (with 2 species) ranges from southern Mexico to 
Venezuela. 


In a three-paper flurry, Bell (1941, 1942a, 1942b) described eight 
new species of what is now Quasimellana, seven of which are valid 
(ricana is a synonym of meridiani Hayward 1984). Bell put all of them 
in genus Atrytone except one: aurora went in Zariaspes, which is no- 
where near Atrytone. Ironically, Bell (1942b:460-465) described aurora 
right after balsa, mexicana, and mulleri, with figures of male genitalia 
detailed enough to show the same (i.e., Quasimellana) kind of cornutus 
in all four species. Although aurora diverged from the other three 
(which are in the eulogius group) in the form of the tegumen, uncus, 


308 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


and gnathos, as well as in the form of the valva, it rather resembled 
nayana, one of the Atrytone species Bell had described the year before. 
Evans (1955) misdetermined his only male of aurora as nayana and 
left aurora in Zariaspes, from which I am finally freeing it. 

Both aurora and nayana came from southwestern Mexico. Steinhau- 
ser (1974) described them again from El Salvador as Mellana tecla and 
M. tamana. 


Quasimellana aurora (Bell, 1942b:464, fig. 7), new combination 
(Figs. 31, 74) 


= tecla (Steinhauser, 1974:15, figs. 41, 42, 81), new synonym, HTé-AME. 


HTé-AMNH. Mexico (Sinaloa, Jalisco, Colima, Guerrero, Oaxaca, PIO EITES)), E] Salvador. 
30 322 (16 629). 

In dorsal view, the tegumen/uncus is narrower and the uncus prongs are closer together 
in aurora than in nayana (Fig. 46), while, in lateral view, the uncus prongs are nearly 
straight (i.e., almost in line with the rest of the tegumen/uncus) in aurora, rather than 
bent sharply upward as they are in nayana (Fig. 47). The valva of aurora, in lateral view, 
has short, blunt, but relatively narrow projections both dorsally and midcaudally, with 
the dorsal projection usually inclined a bit anterodorsally. Both valval projections in nayana 
are broader and more bluntly rounded (the midcaudal projection so much so, that it 
sometimes melts into a totally rounded caudal end, as in Fig. 47). The triangular middle 
process on the inner side of the valva is smaller in aurora than it is in nayana (Fig. 47). 
These two species are superficially very similar, the males with dorsal light areas that are 
more extensive, and usually more orange, than in most other species of Quasimellana, 
but, on an average, the light areas are slightly more extensive in aurora than in nayana 
(compare the aurora and nayana males in figs. 41, 42 and figs. 49, 50, respectively, in 
Steinhauser 1974:30). Two males of aurora from Colima, Mexico, are so extensively light, 
and their orange is so yellow, that they suggest some species of Anatrytone, as. well as 
Quasimellana imperfida. Although both species are medium sized, aurora is a little larger 
than nayana (Fig. 31). 

Genitalia can be deceitful even in published figures. In the lateral views accompanying 
Bell’s original descriptions (Bell 1941:fig. 2, 1942b:fig. 7), the uncus prongs look straight 
not only in aurora (where they should) but also in nayana (where they should bend 
sharply upward). The uncal bend does not show in Bell’s genitalic slide of the nayana 
holotype because the tegumen, uncus, and gnathos are mounted at an odd angle. The 
valva of aurora is poorly drawn in Bell (1942b:fig. 7) but well photographed in Steinhauser 
(1974:fig. 81). Evans’s (1955:pl. 79, fig. M.25.1) caricature of the male genitalia of what 
he calls nayana actually depicts aurora; and Steinhauser’s (1974:figs. 48, 44, 82) photos 
of a tecla (= aurora) female, and her genitalia, really relate to eulogius. 

To match mates can be tricky, especially in strongly dimorphic species. I know from 
comparing series of Quasimellana females that the taxonomically important, midventral, 
posteriorly directed, bristly projection from the posterior margin of the lamella postva- 
ginalis may vary greatly in size and shape among conspecific individuals. It certainly 
varies among the three females I have seen in the nayana subgroup (whose genitalia— 
with their ultrawide, midventral, bristly projection and small lateral lobes [Fig. 74]—are 
immediately distinct). Though all three females may belong to the same species, I am 
tentatively referring the two with shorter bristly projections (Fig. 74) to aurora and the 
one with the bristly projection twice as long (about as in Steinhauser 1974:fig. 84) to 
nayana. Even so, a large measure of individual variation remains since the bristly pro- 
jection is considerably shorter and wider in the unfigured female of aurora than it is in 
the figured female (Fig. 74), and distally truncate, instead of rounded. 


VOLUME 48, NUMBER 4 309 


48 


Fics. 48, 49. Male genitalia of holotype of Quasimellana noka (what remains of 
Evans's dissection), locality unknown, (X-3457) (BMNH). Scale = 1.0 mm. 48, Tegumen, 
uncus, and gnathos in dorsal view. 49, Tegumen, uncus, gnathos, upper vinculum, left 
valva, and distal end of aedeagus, with vesica everted and cornuti stippled, in left lateral 
view (plus primary cornutus in profile). 


Quasimellana nayana (Bell, 1941:1, fig. 2), new combination 
(Figs. ol, 46, 47) 
= tamana (Steinhauser, 1974:18, figs. 49, 50, 83, 84), new synonym, HTé-AME. 


HTé-AMNH. Mexico (Nayarit, Chiapas), Guatemala, El Salvador, Venezuela. 8 6 1 2 
(8 41 9). 
See aurora. 


Ungrouped Species within the sethos Group 
(Figs. 31, 36, 48-57, 75, 76, 93-104) 


This artificial assemblage of 6 species includes an obvious sister pair: 
pazina and antipazina. 


310 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fic. 50. Male genitalia of holotype of Quasimellana pazina (what remains of Evans’s 
dissection) from Yungas and La Paz, 1000 m, BOLIVIA, 1902 (X-3460) (BMNH). Scale 
= 1.0mm. Most of left valva and distal end of aedeagus, with vesica everted and cornutus 
stippled, in left lateral view (plus cornutus in dorsal view). 


Quasimellana noka (Evans, 1955:357, pl. 79, fig. M.25.10), 
new combination 


(Figs. 31, 48, 49, 93, 94) 


HT3-BMNH. No locality. 1 6 (1 4). 

I can safely claim that this genitalically peculiar species, known only from a dataless 
type, comes from somewhere in the neotropics. The valva of noka is unique: two narrow, 
rounded, posterodorsal and midcaudal projections extend not just dorsad and caudad but, 
at the same time, strongly mediad (Fig. 49). The valva lacks a triangular middle process 
on its inner side. Although the uncus is well divided (Fig. 48), the prongs are neither 
widely separated nor somewhat divergent as they are in the nayana subgroup (Fig. 46). 
Instead, they are much as in pazina and antipazina (Fig. 51), with a V-shaped (not U- 
shaped) intervening gap and just a hint of divergence. Since Evans’s dissection wrecked 
the juxta, I cannot say whether it is posterior or anterior in location. Dorsally the light 
areas of the wings are neither extensive and orangy, as in aurora (Steinhauser 1974:fig. 
41), nayana (Steinhauser 1974:fig. 49), sista (Fig. 99), and andersoni (Fig. 101), nor 
reduced and pale yellow, as in inconspicua and angra (Fig. 107). The result is an “average 
look” (Fig. 93), with yellow orange spots—the spotting less developed than in pazina 
(Fig. 95) but much as in myron and verba (Fig. 105). Ventrally a dirty, dull orangish 
shade runs over the hindwing and along the costa and across the apex of the forewing 
(Fig. 94). 


Quasimellana pazina (Evans, 1955:357, pl. 79, fig. M.25.9), 
new combination 


(Figs. 31, 50, 95, 96) 


HTé-BMNH. Bolivia. 2 6 (2 8). 

See antipazina. 

Evans described pazina from a single male. Light areas (which are dorsally yellow 
orange to orange and ventrally yellower) are much more extensive in the second known 
male of pazina than they are in the first (the type, shown in Figs. 95, 96)—so much so, 


VOLUME 48, NUMBER 4 Ole 


Fics. 51,52. Male genitalia of holotype of Quasimellana antipazina from La Estrella, 
1500 m, Cartago, COSTA RICA, 20 May 1979, G. B. Small (X-2592) (USNM). Scale = 
1.0 mm. 51, Tegumen, uncus, and gnathos in dorsal view. 52, Complete genitalia (minus 
right valva), with vesica everted and juxta and cornuti stippled, in left lateral view (plus 
primary cornutus in profile). 


that the second male will not key to pazina in Evans (1955). Dissection of the new male 
reveals that the juxta is anterior in location and confirms the fact that the uncus is divided. 
Evans (1955:pl. 79, fig. M.25.9) clearly showed a divided uncus in his trio of original 
genitalia figures; but the tegumen, uncus, gnathos, and vinculum of his dissected holotype 
have since been lost. 


Quasimellana antipazina, new species 


(Figs. 31, 51, 52, 97, 98) 


HTé6-USNM. Costa Rica. 1 4 (1 8). 

Superficially and dorsally much like myron and verba, with light areas reduced to 
discrete orange yellow spots (Fig. 97); ventral overscaling (narrowly along costa of fore- 
wing, broadly across apex of forewing, and all over the hindwing) not pale, cold, dull 
greenish as in those species but warm, yellowish brown (Fig 98). Light dashes on dorsal 
hindwing distal to cell reduced to only two, in spaces 3 and 4 (Fig. 97)—usually more in 
myron and verba (and other species). Much larger than myron and verba: lone male 
(forewing 17.0 mm) nearly attaining mean male forewing length of mexicana, the giant 
of Quasimellana, and exceeding the larger (16.4 mm) of two known males of pazina 
(Fig. 31). As in pazina, nudum of 15 segments (Table 1) and apiculus exceptionally long 
for Quasimellana (but pazina with better developed light areas dorsally, especially on 
hindwing [Fig. 95], and pazina bright yellow to orange yellow ventrally [Fig. 96], instead 
of yellowish brown). Genitalia (Figs. 51, 52) very like those of pazina (Fig. 50)—including 
juxta anterior in location—but divided uncus narrower, with each prong less massive in 
dorsal view, less elevated distally in lateral view, and proximally lacking ventrolateral 
swelling large enough to show in dorsal view; valva with dorsodistal projection (which 
extends dorsad and mediad) narrower (though with extended, sharp, midcaudal projection 
just as in pazina [compare Figs. 52 and 50]); and tiny, secondary, vestigial cornutus 
present. 


312 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fic. 53. Male genitalia of holotype of Quasimellana sista (what remains of Evans’s 
dissection) from VENEZUELA (X-3459) (BMNH). Scale = 1.0 mm. Most of left valva 
and distal end of aedeagus, with vesica everted and broken cornutus stippled, in left 
lateral view (plus broken cornutus in dorsal view). 


Holotype. 6; COSTA RICA, CARTAGO, La Estrella, 1500 m, 20 May 1979, G. B. 
Small; J. M. Burns genitalia dissection X-2592; USNM. 

Etymology. Quasimellana antipazina is the sister species and counterpart of O. pazina 
in the opposite hemisphere (northern instead of southern). 


Quasimellana sista (Evans, 1955:356, pl. 79, fig. M. = Se 
new combination 
(Figs. 31, 58, 99, 100) 


HTé-BMNH. Venezuela. 1 4 (1 8). 

Placing this species (or determining its sista) is extra difficult because the holotype is 
still the only specimen, Evans (1955:pl. 79, fig. M.25.5) provided just two of his usual 
three genitalic views (omitting the ventral view of the tegumen, uncus, gnathos, and distal 
end of the aedeagus), most of Evans’s dissection of the genitalia is now missing, and the 
parts that remain (valva, aedeagus, cornutus) are broken (Fig. 53). Both the everted vesica 
and what there is of the cornutus clearly indicate the sethos group. For finer placement 
it would help to know whether the juxta is posterior or anterior, and the uncus, divided 
or undivided. Though the (incomplete) valva looks most like that of andersoni (Fig. 55), 
there are significant differences: the sharp, midcaudal projection points dorsad instead of 
caudad, the ventrodistal corner of the valva curves gently instead of angling sharply, and 
the triangular middle process on the inner side has disappeared (Fig. 53). Finally, sista 
lacks the tiny, secondary cornutus; and its big cornutus lacks the ventrally protruding 
sclerotization of andersoni (Fig. 55). Superficially, with extensive light areas and orange 
coloring, sista looks a lot like andersoni, pazina, aurora, and nayana (and it may be 
sympatric with nayana, since both occur in Venezuela—but no further locality data exist 
for sista). However, the orange in space 1b of the dorsal forewing essentially stays in the 
lower half of that space in sista (Fig. 99) while spreading the full height of that space in 
the other four species (Figs. 95, 101 and Steinhauser 1974:figs. 41, 49). 


VOLUME 48, NUMBER 4 | 313 


Fics. 54,55. Male genitalia of paratype of Quasimellana andersoni from Dos Amates, 
Veracruz, MEXICO, 15 January 1972, (X-3004) (Anderson collection). Scale = 1.0 mm. 
54, Tegumen, uncus, and gnathos in dorsal view. 55, Complete genitalia (minus right 
valva), with vesica everted and juxta and cornuti stippled, in left lateral view. 


Quasimellana andersoni, new species 


(Figs. 31, 54, 55, 75, 101-104) 


HTé-USNM. Mexico (Veracruz, Chiapas), Costa Rica. 8 6 2 2 (8 6 2 9). 

Superficially very like aurora and especially nayana of nayana subgroup, with extensive, 
orange, light areas dorsally and ventrally in male and strong sexual dimorphism (Figs. 
101-104). Near nayana in size, probably a little smaller on average (Fig. 31), so one of 
the smallest species of Quasimellana: mean male forewing length and SE = 18.66 + 0.21 
mm, range 12.8-14.5.mm, n = 8; but number of nudum segments (mean 14.8) maximum 
for sethos group (Table 1). Despite posterior juxta (Fig. 55), male genitalia highly distinct 
from nayana subgroup: tegumen/uncus long and narrow, with uncus undivided (Fig. 
54); valva (Fig. 55) perhaps most like sista (qv) (Fig. 53; Evans 1955:pl. 79, fig. M.25.5), 
but also somewhat remindful of myron, verba, pazina (Fig. 50), and antipazina (Fig. 52) 
except for very short, unextended, sharp, midcaudal projection; body of cornutus with 
at least some ventrally protruding sclerotization (Fig. 55) and dorsally more concave than 
in other species of Quasimellana. Midventral, posteriorly directed, bristly projection from 
posterior margin of lamella postvaginalis, delicate (Fig. 75), about as in myron subgroup 
(Fig. 77) and mulleri of eulogius group (Fig. 73); outline of ostium bursae flask-shaped 
in ventral view (Fig. 75); anterior ductus bursae abruptly constricted to half the posterior 
width, in dorsal and ventral view (Fig. 75). 

Holotype. 6; MEXICO, VERACRUZ, Tapalapa, September 1971; J. M. Burns genitalia 
dissection X-3003; USNM. 

Paratypes. n = 7 6 2 2. MEXICO, VERACRUZ: Dos Amates, 15 January 1972, 1 4, J. 
M. Burns genitalia dissection X-3004, collection of R. A. Anderson; Catemaco, December 
1963, 3 6, T. Escalante, H. A. Freeman genitalia dissection H-257, J. M. Burns genitalia 
dissections X-3644, X-3645, AMNH; same data except October 1965, 1 4, S. R. Steinhauser 
genitalia dissection SRS-2702, AME. MEXICO, CHIAPAS, Santa Rosa, Comitan, Septem- 
ber 1963, 1 6 2 2, T. Escalante, H. A. Freeman genitalia dissection H-64, J. M. Burns 
genitalia dissections X-3642, X-3643, AMNH. COSTA RICA, HEREDIA, 3.8 km N Santa 
Clara, 5 September 1987, 1 46, G. & A. Austin, S. R. Steinhauser genitalia dissection SRS- 
2957, collection of G. T. Austin. 


314 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


56 


aS 


Fics. 56, 57. Male genitalia of Quasimellana sethos from Paraiso, Canal Zone, PAN- 
AMA, 27 April 1982, S. S. Nicolay (X-2596) (USNM). Scale = 1.0 mm. 56, Tegumen, 
uncus, and gnathos in dorsal view. 57, Complete genitalia (minus right valva), with vesica 
everted and juxta and cornuti stippled, in left lateral view. 


Etymology. I am pleased to name this species for Richard A. Anderson who generously 
and patiently provided large numbers of Quasimellana from his collection. 


Quasimellana sethos (Mabille, 1889:178, fig. 2), 
new combination 
(Figs. 31, 36, 56, 57, 76) 


Panama, Colombia, Ecuador; Evans (1955) also records a total of three males from 
Nicaragua and Venezuela. 21 6 3 2 (6 6 2 9). 

This, the smallest species of Quasimellana (Fig. 31), is what Godman (1900) called 
Atrytone helva and Evans (1955), Mellana helva. Godman (1900:494), on examining the 
female type of Pamphila helva Méschler (1876) from Surinam and the male type of P. 
sethos Mabille (1889) from Chiriqui (western Panama), erroneously judged them con- 
specific and applied the older name. Godman (1900:pl. 94, figs. 25-29) figured, in color, 
“the type of P. sethos, a male, from Chiriqui, and a female from Coatepec [Veracruz, 
Mexico], agreeing with the type of P. helva,” and, in black and white, the male genitalia. 
The genitalic figure represents this species best (even a Quasimellana-style cornutus 
shows). With orange yellow spots, the figured female looks much like the male; but she 
has well developed subapical spots on the forewing in spaces 6, 7, and 8, where the male 
has only a small point in space 6. The trouble is that this species of Quasimellana (like 
many others) exhibits strong sexual dimorphism in which the spots of the female are 
white instead of yellow to orange, and are less well expressed than those of the male. I 
have carefully studied the original descriptions of Pamphila helva (Méschler 1876), de- 
scribed from a male as well as a female, and P. sethos (Mabille 1889), described from a 
single male. Each description includes critical verbal detail plus a good black and white 
figure of the adult. Without question, sethos is the species long designated as helva; and 
helva is not a species of Quasimellana. 

The saccus of sethos is exceptionally long for Quasimellana—tfrom slightly shorter to 
slightly longer (Fig. 57) than the valva. In lateral outline the valva (Fig. 57):is the simplest 


VOLUME 48, NUMBER 4 | 315 


of the sethos group—about as plain as those of the eulogius group (Figs. 41, 48, 45). The 
body of the cornutus is peripherally wrinkled (Fig. 36)—but much less densely than in 
mulleri (Figs. 37, 38) of the eulogius group. The juxta is anterior and the uncus undivided 
(Figs. 56, 57), a combination of character states that relates to the upcoming myron 
subgroup of four species (qv). The midventral, caudally directed projection from the 
posterior margin of the lamella postvaginalis is just a suggestion of its usual self (Fig. 76). 
Dorsally, in color and pattern, the dimorphic sexes of the diminutive sethos resemble 
those of the larger myron and verba, except that sethos males express a slender spot in 
space 4 usually missing from myron/verba. Ventrally, in fresh specimens of sethos, a 
slight greenish cast—over orange yellow in males and over brownish yellow in females— 
somewhat suggests the dull but more pronounced ventral greenish often visible in the 
myron subgroup. 


The myron Subgroup within the sethos Group 
(Figs. 31, 58-60, 77, 105-108) 


Male genitalia. Juxta: The juxta is anterior in location (Fig. 59). Valva: The valva has 
at least a long, sharp midcaudal projection that points caudad, and often a little mediad 
(Figs. 59, 60). Uncus: The distal end of the uncus is undivided, though usually a bit 
bilobed (Fig. 58). 

Female genitalia. The midventral, posteriorly directed, bristly projection from the 
posterior margin of the lamella postvaginalis is delicate (Fig. 77), about as in andersoni 
(Fig. 75) and mulleri (Fig. 73). The posterior part of the sclerotized ductus bursae usually 
extends farther caudad, bringing the ostium bursae closer to the midventral, bristly 
projection (Fig. 77). The anterior part of the sclerotized ductus bursae has a central, 
longitudinal, unsclerotized zone (Fig. 77). 

General features. Male spotting varies from average to much reduced, with many 
males dark enough that the sexual dimorphism typical of Quasimellana, though present, 
is not striking. Ventral overscaling in both sexes often gives a pale, dull greenish cast. The 
species are medium sized to small (Fig. 31). 

Distribution. The myron subgroup (with 4 species) ranges from Mexico to Bolivia and 
Brazil. 


Evans (1955) treated three of the four taxa in this subgroup (myron, 
verba, and inconspicua) as subspecies of myron, describing the fourth 
(angra) as a separate species. Though all four are closely related, male 
genitalia show conclusively that two pairs of sister taxa are involved: 
myron and verba, on the one hand, and inconspicua and angra, on the 
other. The distal end of the valva in myron and verba has a major 
dorsal projection—totally lacking in inconspicua and angra (Figs. 59, 
60)—which extends dorsad and mediad so that the valva as a whole 
resembles that of pazina and antipazina (Figs. 50, 52). Also, the dorsal 
process on the inner surface of the valva is long and conspicuous (as in 
most members of the sethos group) in myron and verba but short and 
inconspicuous in inconspicua and angra (Figs. 59, 60). Female genitalia 
support this grouping, though less showily: the posterior part of the 
ductus bursae is much longer than the anterior part in myron and verba 
(Fig. 77) but only a little longer in inconspicua and angra. 


316 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


58 


Fics. 58,59. Male genitalia of Quasimellana inconspicua from Puyo, 1000 m, Pastaza, 
ECUADOR, 19 October 1989, S. S. Nicolay (X-2826) (USNM). Scale = 1.0 mm. 58, 
Tegumen, uncus, and gnathos in dorsal view. 59, Complete genitalia (minus right valva), 
with vesica everted and juxta and cornutus stippled, in left lateral view. 


(I will say nothing further about females within the myron subgroup 
because sound interspecific differences in genitalia and external ap- 
pearance are not evident among the few specimens at hand.) 

Although males of myron and verba are not difficult to recognize as 
a unit, they are hard to separate from each other. Evans (1955) described 
verba (from Costa Rica, Colombia, Ecuador, and Guyana) as a southern 
subspecies of myron (from Mexico, Nicaragua, and Costa Rica). But 
distinguishing characters neither hold completely nor neatly divide 
geographically. All things considered, these skippers look more like 
overlapping species than subspecies. A quick and easy alternative would 
be a single, too variable, species, at least as dissatisfying as eulogius 
(and unfortunately, for supplemental analysis, far rarer in collections). 
The myron/verba complex now appears to range from central Mexico 
to southern Brazil. 

By contrast, inconspicua and angra are strictly South American, with 
inconspicua western (Ecuador to Bolivia) and angra to the east. Though 
their genitalia look the same and their wings look similarly dark, they 
differ enough in color, pattern, and size (Fig. 31) to warrant specific 
rank. Since Evans’s (1955:358) lists of specimens examined include an 
inconspicua male from “Chanchamayo,” Peru, and an angra male from 
‘“Perene,” Peru, since both names refer to the same general collecting 
area, and since both skippers live at low elevation (even the western 
inconspicua is known only from 250-1000 m), inconspicua and angra 
may well be in contact or sympatric. 


VOLUME 48, NUMBER 4 317 


Fic. 60. Male genitalia of holotype of Quasimellana angra (what remains of Evans's 
dissection) from Para, BRAZIL, H. W. Bates (X-3461) (BMNH). Scale = 1.0 mm. Left 
valva and aedeagus, with vesica everted and cornuti stippled, in left lateral view (plus 
primary cornutus in dorsal view). 


Quasimellana myron (Godman, 1900:498, pl. 94, figs. 20-24), 
new combination 
(Figs. 31, 77) 


Mexico (San Luis Potosi, Veracruz, Guerrero, Oaxaca, Quintana Roo), Guatemala, 
Honduras, Nicaragua, Costa Rica, Panama. 16 652 (9 6 4 Q). 

The dorsodistal projection of the valva is broader and distally rounder than it is in 
verba, and the valva has no middle process on its inner surface (see Evans 1955:pl. 79, 
fig. M.25.12 myron). The secondary, vestigial cornutus is usually absent (present in two 
males from Oaxaca, Mexico, but smaller than it is in verba). Male spot development in 
myron and verba is generally about average to somewhat reduced (Figs. 105, 106); but, 
in the specimens at hand (as in any appreciable series of Quasimellana), it varies con- 
siderably. Evans’s (1955) observation that the yellow dorsal markings of the male are 
paler in myron and darker in verba appears valid but ever so subtle. I find the yellow 
spot in space 2 of the dorsal forewing more helpful: it tends to be rectangular (often 
squarish), with its inner edge more or less vertical, in males of myron but more nearly 
triangular (and wider), with its inner edge oblique, in males of verba (Fig. 105). 


Quasimellana verba (Evans, 1955:358, pl. 79, fig. M.25.12), 
new combination 
(Figs. 31, 105, 106) 


HTé-BMNH. Mexico (San Luis Potosi, Yucatan), Panama, Colombia, Venezuela, Ec- 
uador, Brazil (Sao Paulo). 12 63 ¢ (12 61 9). 

The dorsodistal projection of the valva is narrower and distally more pointed than it 
is in myron, and the valva usually has a middle process on its inner surface (see Godman 
1900:pl. 94, fig. 22 and Evans 1955:pl. 79, fig. M.25.12 verba)—but that process is small 
in three males from Mexico and Panama and absent in two males from Ecuador and 
Brazil. The secondary, vestigial cornutus is usually present (absent in the male from 


318 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Brazil). See myron for slight average differences between myron and verba in the external 
appearance of males. 


Quasimellana inconspicua (Hayward, 1950a:465, fig. 5), 
new combination 
(Figs. 31, 58, 59) 


HTé-IML. Ecuador, Peru, Bolivia. 8 6 2 2 (8 6 1 9). 

Spotting is so reduced in males of both inconspicua and angra that they look mostly 
dark. Their dorsal forewing bears a total of three small yellow spots in the lower part of 
space 1b and in spaces 2 and 3. Even though inconspicua is the larger species, averaging 
more than a millimeter longer than angra in wing length (Fig. 31), its spots are smaller 
and relatively diffuse—much less sharply defined than they are in angra (Fig. 107). 
Moreover, in inconspicua the yellow of these spots is a little paler; and the spot in space 
3 is slightly distad of the spot in space 2, whereas in angra the spot in space 3 slightly 
overlaps the spot in space 2 (Fig. 107). Ventrally, males of inconspicua are darker and 
vaguely greenish brown when fresh, whereas angra males are lighter and somewhat 
yellowish brown. For whatever it may be worth with such small numbers, I note that the 
secondary, vestigial cornutus is absent in nearly half the dissected males of inconspicua 
(Fig. 59) but present in all four dissected males of angra (Fig. 60). 

Specimens of inconspicua have been misdetermined as pandora, a superficially similar 
species or subspecies of Quasimellana in the nicomedes group and a very close sister of 
meridiani. See meridiani for a discussion of external differences between inconspicua 
and pandora. 


Quasimellana angra (Evans, 1955:358, pl. 79, fig. M.25.13), 
new combination 
(Figs. 31, 60, 107, 108) 


HTé-BMNH. French Guiana, Brazil (Para, Maranhao, Pernambuco, Bahia, Minas Ger- 
ais, Rondénia); Evans (1955) also lists one male each from Venezuela, Ecuador, and Peru. 
6342 (4 62 2). 

See inconspicua. 

This is the second smallest species of Quasimellana, exceeding only sethos (Fig. 31). 


The nicomedes Group 
(Figs. 31, 34, 35, 39, 61-67, 78-82, 109-112) 


Male genitalia. Juxta: As in some members of the sethos group, the juxta is anterior 
in location (Figs. 62, 65, 67); but the level of the twist in each lateral arm of the juxta is 
high (Figs. 39, 62, 65, 67), not low as it is in all other species of Quasimellana (Figs. 41, 
43, 45, 47, 52, 55, 57, 59). Cornutus: The head-and-neck of the cornutus is short; it rises 
almost vertically from the body of the cornutus (Figs. 34, 35, 62, 65, 67). No member of 
the nicomedes group has the tiny, secondary, vestigial cornutus that usually occurs in 
other species of Quasimellana. Vesica: The everted vesica is relatively short, fat, and 
irregular (Figs. 62, 65, 67). Valva: The valva has neither a middle process nor a dorsal 
process on its inner surface (Figs. 62, 65, 67) (one or both processes are present in other 
species of Quasimellana). Distally the valva terminates in one or two dorsally directed, 
sharp points (Figs. 65, 67), except in fieldi (Fig. 62). Uncus: In lateral view, the distal 
end of the uncus is much thicker than the distal end of the gnathos (Figs. 62, 65, 67), 
except in amicus where (as in all other species of Quasimellana) the uncus is only a little 
thicker than the gnathos. Again, in lateral view, the uncus is usually (but not always) a 
little shorter than the gnathos (Figs. 62, 65, 67), except in amicus. Tegumen: In dorsal 
view the tegumen generally tapers more abruptly to the uncus (Figs. 61, 63, 64, 66) than 
it does in other species. 


VOLUME 48, NUMBER 4 319 


Fics. 61, 62. Male genitalia of Quasimellana fieldi from San Salvador, EL SALVA- 
DOR, 29 December 1953, M. Salazar (X-2589) (USNM). Scale = 1.0 mm. 61, Tegumen, 
uncus, and gnathos in dorsal view. 62, Complete genitalia (minus right valva), with vesica 
everted and juxta and cornutus stippled, in left lateral view. 


Female genitalia. The ductus bursae extends caudad ventrally but not laterally to form 
a unique scoop beneath the ostium bursae (Figs. 78-82). At its anterior end the sclerotized 
ductus bursae is longitudinally grooved and more or less expanded (Figs. 78-82). The 
anterior apophyses are neither long (as in the eulogius group) nor short (as in the sethos 
group). The only species outside of the nicomedes group with anterior apophyses of 
intermediate length are sethos and (in some individuals) balsa. The corpus bursae is 
relatively short and fat (Figs. 78-82), though less so in fieldi. 

General features. Sexual dimorphism runs the gamut from nonexistent (amicus), through 
weak (nicomedes, meridiani, and presumably pandora), to strong (imperfida, mielkei, 
and fieldi); color pattern, from mostly dark (meridiani and especially pandora), through 
roughly half-and-half (amicus and nicomedes), to mostly light (in the predominantly 
yellow orange males of fieldi, mielkei, and especially imperfida). The species are small 
to medium in size (Fig. 31). 

Distribution. Although the nicomedes group (with 7 species) extends from Mexico to 
Argentina, it is mainly South American. 


Quasimellana amicus (Bell, 1942a:2, fig. 3), new combination 
(Figs. 31, 78) 


HT38-AMNH. Ecuador. 8 6 4 2 (5 6 8 2). 
Superficially this smallish species (Fig. 31) seems out of place in Quasimellana because 
females look just like males, the discal spotband (yellow orange) runs boldly and contin- 


320 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


uously (cut by dark veins) up the forewing from space lb to space 6 (or rarely 7) without 
the usual break in space 5 (distal to the upper half of the cell), and the ventral hindwing 
flaunts a dark border (interrupted by yellow in space lc). This is the sole member of the 
nicomedes group in which the uncus is undivided and (in lateral view) only a little thicker 
than the gnathos. The long, low valva ends distally in a single, dorsally (and medially) 
directed, sharp point. The midventral, posteriorly directed, bristly projection from the 
posterior margin of the lamella postvaginalis is exceptionally delicate and long (extending 
far caudad of the paired, sclerotized, lateral lobes that flank it); its base is not quite hidden, 
in ventral view, by the posterior edge of the ventral scoop of the ductus bursae; and the 
longitudinally grooved, anterior end of the sclerotized ductus bursae is diagonal, slanting 
sharply forward (at about 45°) from right to left (Fig. 78). 


Quasimellana fieldi (Bell, 1942a:2, fig. 4), new combination 
(Figs. 31, 34, 89, 61, 62) 


= montezuma (Freeman, 1969:41, pl. 13, figs. 5-8; pl. 15, fig. 10), new synonym, HTé- 
AMNH. 


HTé-USNM. Mexico (San Luis Potosi), Guatemala, El Salvador, Nicaragua; Evans (1955) 
gives Costa Rica. 54 613 2 (10 638 9). 

The only North American species of the nicomedes group, fieldi superficially resembles 
members of the eulogius group (especially mexicana, eulogius, and balsa) inhabiting the 
same general region. The long, low valva comes dorsodistally to a blunt or vaguely pointed 
end (Fig. 62)—not a sharp point as in amicus. The uncus prongs (in dorsal view) are 
short, parallel, and more or less rounded at their distal ends (Fig. 61). The body of the 
cornutus is large and mostly flat (Figs. 34, 62). The midventral, posteriorly directed, bristly 
projection from the posterior margin of the lamella postvaginalis is medium in length 
(extending a little caudad of the paired, sclerotized, lateral lobes that flank it) and about 
as wide as it is in mielkei (Fig. 81) or meridiani (Fig. 82) to considerably wider. The 
longitudinally grooved, anterior end of the sclerotized ductus bursae is diagonal, much 
as it is in amicus (Fig. 78), but it slants forward from right to left even more steeply than 
in that species. (See Steinhauser 1974:fig. 87 for a photograph of the female genitalia of 
fieldi in ventral view.) 

When Freeman (1969) described montezuma (which is synonymous with fieldi), he 
grouped it with what he called its “nearest relatives’ —nayana and mulleri—in a “nayana 
complex,” based only on a few similarities in external appearance. In reality, these three 
species are not even close, each belonging in a different one of the three species groups 
of Quasimellana. 


Ouasimellana nicomedes (Mabille, 1883:LXX), new combination 
(Figs. 31, 35, 79, 80) 


= monica (Plotz, 1886:98), new synonym, HT?-ZMHB. 


HT8-ZMHB. Brazil (Parana, Santa Catarina); Evans (1955) gives localities to the north 
in the neighboring states of Sao Paulo and Rio de Janeiro. 46 42(363 9). 

In the modest sexual dimorphism of nicomedes, the female expresses spots almost as 
well as the male, and her spots are a malelike yellow orange (not whitish and semihyaline 
on the forewing as they are in most species of Quasimellana). The long, low valva comes 
dorsodistally either to a single, sharp, dorsally directed point, as in amicus, or to two such 
points, as in imperfida, mielkei (Fig. 65), pandora (Fig. 67), and meridiani—but in 
nicomedes the points are a little closer together, and the distal point is higher than the 
proximal point instead of the other way around. The uncus prongs (in dorsal view) are 
very short, barely divergent, and truncate at their distal ends. The body of the cornutus 
is narrow and highly convex (Fig. 35). The midventral, posteriorly directed, bristly 
projection from the posterior margin of the lamella postvaginalis (Figs. 79, 80) is delicate 
and long, extending somewhat caudad of the paired, sclerotized, lateral lobes that flank 
it—but not as far as in amicus (Fig. 78). 


VOLUME 48, NUMBER 4 321 


Quasimellana imperfida, new species 


(Figs. 31, 63) 


HTé-USNM. Brazil (Mato Grosso, Goias), Bolivia; Evans (1955) also lists Colombia and 
Paraguay under the name Mellana perfida. 4622 (462 9). 

I have scrutinized the type (¢-ZMHB), its genitalia (X-3763), and the original description 
of perfida Moschler (1878:221) and found that it is a species of Anatrytone and that what 
everyone is currently calling Mellana perfida is an undescribed species of Quasimellana. 

See mielkei. Size similar to mielkei, maybe a little smaller (Fig. 31): mean male forewing 
length 15.4 mm, range 14.8-16.2 mm, n = 4; mean female forewing length 16.8 mm, 
range 16.5-17.0 mm, n = 2. Nudum 14 or, usually, 15 segments (Table 1). 

Externally, the light males of imperfida suggest males of Anatrytone—especially A. 
perfida or some combination of that species and A. logan. Quasimellana imperfida is a 
close sister to mielkei (qv); and these two are related, on the one hand, to the sisters 
meridiani/pandora and, on the other, to nicomedes (qv). The tegumen, uncus, and 
gnathos of imperfida (Fig. 63) are about as in nicomedes, while the cornutus is about as 
in meridiani and pandora (Fig. 67), i.e., with the body slightly less narrow and rather 
less highly convex (flatter) than in nicomedes. In lateral view the valva is not quite as 
low as in fieldi (Fig. 62) and especially nicomedes and amicus; in this regard, it is much 
like that of meridiani and pandora (Fig. 67) but longer. The female genitalia of imperfida 
are similar to those of meridiani (Fig. 82) and presumably pandora. 

Holotype. 6; BRAZIL, MATO GROSSO, Colegio Buriti, Chap. Guimaraes, 700 m, 26 
May 1969, S. S. Nicolay; S. S. Nicolay genitalia dissection H435 (left valva missing); 
USNM. 

Paratypes. n = 3 6 2 2. BRAZIL: MATO GROSSO: Nivac, 1 6, J. M. Burns genitalia 
dissection X-3462, collection of O. H. H. Mielke; Buriti, Chapada dos Guimaraes, 600 m, 
27 June 1972, 1 2, Mielke & Brown, X-3463, specimen number DZ 3519 in UFPR; Caceres, 
7-9 February 1985, 1 6, C. Elias, X-3116, DZ 3513 in UFPR; GOIAS, Goias Velho, 25 
February 1979, 1 9, Gifford, X-3464, DZ 3521 in UFPR. BOLIVIA, Chiquitas, 1 4, X-3764, 
ZMHB. 

Etymology. Quasimellana imperfida simply is not perfida. Ever since Evans (1955), 
it has been going under the name Mellana perfida; but perfida is really a superficially 
similar looking species of Anatrytone (see p. 286). 


Quasimellana mielkei, new species 


(Figs. 31, 64, 65, 81, 109-112) 


HTé-UFPR. Brazil (Minas Gerais). 1 612 (161 9). 

Close sister to imperfida. Sexual dimorphism strong (Figs. 109, 111), as in imperfida 
and fieldi. Wings fuller than in imperfida. Facies much as in imperfida, but dorsally, in 
male, extensive light areas of wings more orange (imperfida yellower), and transition 
between broad, light areas and narrow, dark, outer borders less hard and sharp; dorsally, 
in female, semihyaline spot in cell of forewing double (Fig. 111) instead of single as in 
imperfida (with so little material, this difference perhaps nothing more than individual 
variation); ventrally, in both sexes, dark ground color evident through thin, light over- 
scaling (Figs. 110, 112)—wings ventrally not almost all bright yellow as in imperfida. 
Genitalia in both sexes (Figs. 64, 65, 81) like those of imperfida (including small, flexible, 
well sclerotized, conspicuous, and coarsely spinulose scouring-pad cornutus in male [Fig. 
65], not present in other species of Quasimellana) except uncus slightly wider in dorsal 
view (compare Figs. 64 and 63), aedeagus somewhat stouter, and ductus bursae slightly 
wider in ventral or dorsal view. Size similar to imperfida, maybe a little larger (Fig. 31): 
forewing length of male 16.0 mm; of female 17.0 mm. Nudum long—16 segments in 
both specimens. 

Holotype. 6; BRAZIL, MINAS GERAIS, Diamantino, November 1977, Sakakibare; J. 
M. Burns genitalia dissection X-3119; specimen number DZ 3515 in UFPR. 

Paratype. °; same data; X-3120; DZ 3514 in UFPR. 

Etymology. I am delighted to name this species for Olaf H. H. Mielke who has 
enormously increased our knowledge of neotropical skippers and has generously lent 
much helpful material. 


322 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 63-65. Male genitalia of paratype of Quasimellana imperfida from Caceres, 
Mato Grosso, BRAZIL, 7-9 February 1985, C. Elias (X-3116) (UFPR) and ‘of holotype of 
Q. mielkei from Diamantino, Minas Gerais, BRAZIL, November 1977, Sakakibare (X- 
3119) (UFPR). Scale = 1.0 mm. 63, Tegumen, uncus, and gnathos of imperfida in dorsal 
view. 64, Tegumen, uncus, and gnathos of mielkei in dorsal view. 65, Complete genitalia 
of mielkei (minus right valva), with vesica everted and juxta and cornuti stippled, in left 
lateral view. 


Quasimellana meridiani (Hayward, 1934:117, pl. 5, fig. 16), 
new combination 
(Figs. 31, 82) 


= ricana (Bell, 1941:1, fig. 1), HTé-AMNH. 


Brazil (Parana), Paraguay, Argentina (Misiones). 4 6 1 2 (3 6 1 9). 

Though genitalically close to the sisters imperfida and mielkei, sisters meridiani and 
pandora depart widely from them superficially: wings are relatively elongate and narrow 
(much as in various species of Panoquina), and are mostly blackish brown above. Males 
of meridiani are dorsally darkened enough to resemble females of imperfida, while males 
of pandora—their forewings usually with small yellow orange spots only in spaces 1b, 2, 
and 3—are so dark that they look like males of inconspicua of the sethos group. (However, 
spots in pandora are more orange than they are in inconspicua; the spot in space 3 is 
more distad of the spot in space 2 in pandora than it is in inconspicua; and the spot in 


VOLUME 48, NUMBER 4 . 323 


Fics. 66, 67. Male genitalia of Quasimellana pandora from Rio Xingu Camp, 52° 
22’ W, 3° 39’ S (first jungle stream trail 1, malaise trap day and night collection), ca 60 
km S Altamira, Para, BRAZIL, 2-8 October 1986, P. Spangler and O. Flint (X-2673) 
(USNM). Scale = 1.0 mm. 66, Tegumen, uncus, and gnathos in dorsal view. 67, Complete 
genitalia (minus right valva), with vesica everted and juxta and cornutus stippled, in left 
lateral view (plus cornutus in profile). 


space lb is, in pandora, often a sort of V on its side, spanning the height of space 1b, 
instead of a mere dot or bar, always limited to the lower part of that space, as it is in 
inconspicua. Moreover, males of meridiani and pandora average about a millimeter 
longer in the forewing than do males of inconspicua [Fig. 31]). Spotting in the female of 
meridiani is reduced from that in the male, but the spots are still a pale yellow orange; 
the female of pandora is unknown (spots in the female of inconspicua are whitish). 
Ventrally males of meridiani are mostly yellow but duller than in imperfida, and males 
of pandora are duller still (especially in Ecuador). The uncus prongs of meridiani and 
pandora are higher and wider than they are in other members of the nicomedes group: 
in left lateral view, the rounded, ventral lobe of the left prong projects exceptionally far 
downward (Fig. 67); in dorsal view, the prongs are truncate but distinctly divergent, 
usually with a good, triangular notch between them (Fig. 66). The valva (Fig. 67) is about 
as in imperfida and mielkei (Fig. 65) but shorter. The female genitalia (Fig. 82) are 
similar to those of imperfida and mielkei (Fig. 81), but the midventral, posteriorly directed, 
bristly projection from the posterior margin of the lamella postvaginalis is a little heavier 
and wider, at least in meridiani (the female of pandora is unknown). 


Quasimellana pandora (Hayward, 1940:867, fig. 20), 
new combination 
(Figs. 31, 66, 67) 


HT6-IML. Ecuador, Peru, Bolivia, French Guiana, Brazil (Para, Distrito Federal); Evans 
(1955) also lists Guyana. 14 6 (18 8). 

See meridiani. 

Evans (1955) may have been correct in treating meridiani and pandora as subspecies. 
More material is needed to resolve this matter. 


324 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fics. 68, 69. Female genitalia of Quasimellana mexicana from Rio Metlec, Fortin 
de las Flores, Veracruz, MEXICO, 2 October 1975, J. Powell and J. Chemsak (X-2809) 
(UCB). Scale = 1.0 mm. 68, Sterigma, bursa copulatrix, and an indication of the terminal 
abdominal tergites (including the anterior and posterior apophyses and the ovipositor 
lobes) in ventral view. 69, The same, plus part of the ductus seminalis, in right lateral 
v1lew. 


VOLUME 48, NUMBER 4 325 


Fics. 70, 71. Female genitalia of Quasimellana balsa from Rte. 16, km 242-254, 
Sonora, MEXICO, 1 September 1991, J. P. Brock (X-3656) (USNM). Scale = 1.0 mm. 70, 
Sterigma, bursa copulatrix, and an indication of the terminal abdominal tergites (including 
the anterior and posterior apophyses and the ovipositor lobes) in ventral view. 71, The 
same, plus part of the ductus seminalis, in right lateral view. 


326 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 72, 73. Female genitalia of Quasimellana showing the sterigma, bursa copu- 
latrix, and an indication of the terminal abdominal tergites (including the anterior and 
posterior apophyses and the ovipositor lobes) in ventral view. Scale = 1.0 mm. 72, Q. 
eulogius from Cayuga, GUATEMALA, September (X-3062) (USNM). 73, Q. mulleri 
from Victoria, Tamaulipas, MEXICO, 16 August 1962, Stallings, Turner (X-3648) (AMNH). 


VOLUME 48, NUMBER 4 | SPATl 


Fics. 74, 75. Female genitalia of Quasimellana showing the sterigma, bursa copu- 
latrix, and an indication of the terminal abdominal tergites (including the anterior and 
posterior apophyses and the ovipositor lobes) in ventral view. Scale = 1.0 mm. 74, Q. 
aurora from Las Juntas Verano, 1000 ft (305 m), Hwy. 200 below Puerto Vallarta, Jalisco, 
MEXICO, 8 August 1989, J. Kemner (X-2823) (USNM). 75, Q. andersoni (paratype) 


from Santa Rosa, Comitan, Chiapas, MEXICO, September 1963, T. Escalante (X-3643) 
(AMNH). 


328 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Fics. 76, 77. Female genitalia of Quasimellana showing the sterigma and bursa 
copulatrix in ventral view. Scale = 1.0 mm. 76, Q. sethos, with an indication of the 
terminal abdominal tergites (including the anterior and posterior apophyses and the 
ovipositor lobes) and the ductus seminalis, from Howard Air Force Base, Canal Area, 
PANAMA, 3 February 1985, R. A. Anderson (X-3006) (USNM). 77, Q. myron from 
Coatepec, Veracruz, MEXICO, (X-2600) (USNM). 


VOLUME 48, NUMBER 4 329 


Fic. 78. Female genitalia of Quasimellana amicus from Napac, 1000 m, Pichincha, 
ECUADOR, 28 September 1975, S. S. Nicolay (X-2852) (USNM). Scale = 1.0 mm. Ste- 
rigma, bursa copulatrix, and an indication of the terminal abdominal tergites (including 
the anterior and posterior apophyses and the ovipositor lobes) in ventral view. 


330 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 79, 80. Female genitalia of Quasimellana nicomedes from Cacatu, Antonina, 
20 m, Parana, BRAZIL, 25 April 1973, Mielke (X-2367) (MacNeill collection). Scale = 
1.0 mm. 79, Sterigma, bursa copulatrix, and an indication of the terminal abdominal 
tergites (including the anterior and posterior apophyses and the ovipositor lobes) in ventral 
view. 80, The same, plus part of the ductus seminalis, in right lateral view. 


VOLUME 48, NUMBER 4 ool 


Fics. 81, 82. Female genitalia of Quasimellana showing the sterigma, bursa copu- 
latrix, and an indication of the terminal abdominal tergites (including the anterior and 
posterior apophyses and the ovipositor lobes) in ventral view. Scale = 1.0 mm. 81, Q. 
mielkei (paratype) from Diamantino, Minas Gerais, BRAZIL, November 1977, Sakakibare 
(X-3120) (UFPR). 82, OQ. meridiani from Dos de Mayo, Misiones, ARGENTINA, 7 March 
1989, Foerster (X-3614) (Mielke collection). 


332 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 83-97. Mostly holotypes (HT) and paratypes (PT) of Anatrytone and Quasi- 
mellana in dorsal (D) and (&) ventral (V) views (all x1). 83, 84, Anatrytone sarah 4, 
HT, D & V, Limoncocha, 240 m, Napo, ECUADOR (S. S. Nicolay genitalia dissection 
H739) (USNM). 85, 86, A. sarah 2°, PT, D & V, Puerto Maldonado, 290 m, PERU (X- 
2607) (USNM). 87, 88, Quasimellana mexicana 8, HT, D & V, La Gloria, Cardel, 
Veracruz, MEXICO (W. D. Field genitalia dissection 1820) (USNM). 89, 90, QO. mexicana 
2, D & V, Rio Metlec, Fortin de las Flores, Veracruz, MEXICO (X-2809) (UCB). 91, 92, 
Q. siblinga 6, HT, D & V, El Vado-San Sebastian, 1675-1980 m, Oaxaca, MEXICO 
(USNM). 93, 94, QO. noka 6, HT, D & V, locality unknown (X-3457) (BMNH). 95, 96, 
QO. pazina 6, HT, D & V, Yungas and La Paz, 1000 m, BOLIVIA (X-3460) (BMNH). 97, 
Q. antipazina 8, HT, D, La Estrella, 1500 m, Cartago, COSTA RICA (X-2592) (USNM). 


VOLUME 48, NUMBER 4 333 


Fics. 98-112. Holotypes (HT) and paratypes (PT) of Quasimellana in dorsal (D) and 
(&) ventral (V) views (all x1). 98, Q. antipazina 6, HT, V, La Estrella, 1500 m, Cartago, 
COSTA RICA (X-2592) (USNM). 99, 100, QO. sista 6, HT, D & V, VENEZUELA (X- 
3459) (BMNH). 101, 102, Q. andersoni 6, HT, D & V, Tapalapa, Veracruz, MEXICO 
(X-3003) (USNM). 103, 104, QO. andersoni 2, PT, D & V, Santa Rosa, Comitan, Chiapas, 
MEXICO (X-3643) (AMNH). 105, 106, QO. verba 6, HT, D & V, Balzapamba, Bolivar, 
ECUADOR (X-3458) (BMNH). 107, 108, Q. angra 6, HT, D & V, Para, BRAZIL (X- 
3461) (BMNH). 109, 110, QO. mielkei 6, HT, D & V, Diamantino, Minas Gerais, BRAZIL 
(X-3119) (UFPR). 111, 112, O. mielkei?, PT, D & V, Diamantino, Minas Gerais, BRAZIL 
(X-3120) (UFPR). 


334 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


ACKNOWLEDGMENTS 


Museums providing material appear below, with abbreviations (when cited above) and 
the names of workers who kindly arranged loans: Allyn Museum of Entomology, Florida 
Museum of Natural History, Sarasota, Florida (AME), Stephen R. Steinhauser; American 
Museum of Natural History, New York, New York (AMNH), Frederick H. Rindge; 
Carnegie Museum of Natural History, Pittsburgh, Pennsylvania (CMNH), John E. Raw- 
lins; Departamento de Zoologia, Universidade Federal do Parana, Curitiba, Parana, Brazil 
(UFPR), Olaf H. H. Mielke; Essig Museum of Entomology, University of California, 
Berkeley, California (UCB), Jerry A. Powell; Fundacién e Instituto Miguel Lillo, Univ- 
ersidad Nacional de Tucuman, San Miguel de Tucuman, Argentina (IML), Zine A. de 
Toledo and A. Willink; Milwaukee Public Museum, Milwaukee, Wisconsin, Allen M. 
Young; Museo de Historia Natural, Universidad Nacional Mayor de San Marcos, Lima, 
Peru (MUSM), Gerardo Lamas; Museum ftir Naturkunde der Humboldt-Universitat zu 
Berlin, Zoologisches Museum, Berlin, Germany (ZMHB), H. J. Hannemann and W. Mey; 
National Museum of Natural History, Smithsonian Institution, Washington, District of 
Columbia (USNM); Natural History Museum, London, England (BMNH), Philip R. Ack- 
ery. 

The following people lent and/or donated material from their collections: Richard A. 
Anderson, Jim P. Brock, H. Avery Freeman, Jeffrey Glassberg, Bernard Hermier, Daniel 
H. Janzen, John Kemner, Dan L. Lindsley, C. Don MacNeill, Olaf H. H. Mielke, Doug 
Mullins, S. S. Nicolay, John A. Shuey, and J. Bolling Sullivan. Warren Steiner spread some 
of it. The late Adrienne Venables and especially Elizabeth Klafter dissected several 
hundred genitalia, a selection of which Young Sohn drew. Victor E. Krantz photographed 
adults plus the Godman genitalic figure. Jerry A. Louton calculated statistics on size and 
prepared Fig. 31, which George L. Venable polished. Don Harvey, Bernard Hermier, 
Gerardo Lamas, and Robert Robbins furnished useful information. Sarah Burns helped 
again and again in various ways. George T. Austin and C. Don MacNeill reviewed the 
manuscript. The Biodiversity Programs, National Museum of Natural History, Smithson- 
ian Institution, provided funds for trips to two museums. Shapiro, Bernstein & Co., Inc., 
New York, New York, publisher (January 1923) of the song “Yes! We Have No Bananas, ” 
permitted me to borrow from its title in exchange for a reprint of this paper. Thank you 
one and all. 


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Received for publication 4 May 1994; accepted 19 June 1994. 


Journal of the Lepidopterists’ Society 
48(4), 1994, 338-372 


IMMATURE STAGES OF VENEZUELAN DIOPTINAE 
(NOTODONTIDAE) IN JOSIA AND THIRMIDA 


JAMES S. MILLER 


Department of Entomology, American Museum of Natural History, 
Central Park West at 79th Street, New York, New York 10024, USA 


AND 


L. DANIEL OTERO 


Laboratorio de Quimica Ecolégica, Departamento de Quimica, Facultad de Ciencias, 
Universidad de Los Andes, La Hechicera, Mérida 5101, Venezuela 


ABSTRACT. Immature stages of five species of Dioptinae (Notodontidae) from Ven- 
ezuela are described and illustrated. Four are in the genus Josia Hubner and one is in 
Thirmida Walker. All feed as larvae on Passiflora (Passifloraceae). This is the first life 
history reported for a member of Thirmida. Immature stages are described for each 
species, and distributional and hostplant data are provided. The tribe Josiini, formerly a 
subfamily (Josiinae), is delineated. It comprises 103 species in eleven dioptine genera. 
Josia turgida and Thirmida discinota, treated by previous authors as subspecies, are 
revised to species status. 


Additional key words: Josiini, Passiflora, montane forest, larval morphology. 


The Neotropical moth subfamily Dioptinae is unique among Noto- 
dontidae in containing many diurnal forms with aposematic color pat- 
terns (Hering 1925, Kohler 19380, Miller 1992a). The group includes 
over 400 described species (Byrk 1930), most of which are poorly rep- 
resented in museums worldwide. The most commonly collected diop- 
tines are those in Josia, a genus distributed from southern Mexico south 
to northern Argentina. Josia adults are strikingly colored, usually with 
orange or yellow markings against a dark ground color, and they tend 
to fly in a slow, fluttering manner. The caterpillars, which feed on plants 
in the genus Passiflora (Passifloraceae), commonly called “passion flow- 
ers,” are conspicuously patterned with various hues of reddish maroon, 
yellow, and white. 

Among Lepidoptera only two large species radiations are associated 
with Passiflora—Josia and its relatives, and members of the butterfly 
subfamily Heliconiinae (Nymphalidae). Heliconius, containing ap- 
proximately 70 Neotropical species (Brown 1981, DeVries 1987), has 
been the subject of intense ecological and evolutionary research, and 
an immense amount of biological data has accumulated. For example, 
there are over 350 published host records for the genus (Benson 1978). 
In contrast, the amount of biological information available for Josia is 
minute; hosts are recorded for only five species, all from Brazil (Spitz 
1931, d’Almeida 1932, Bienzanko 1962a, 1962b, Silva et al. 1968). In 
this paper we describe the immature stages of five Passiflora-feeding 
Dioptinae from Venezuela. 


VOLUME 48, NUMBER 4 339 


TABLE 1. Genera belonging in the tribe Josiini (Notodontidae: Dioptinae) based on 
presence of a kettle-drum metathoracic tympanum in the adult (see text). Generic names 
follow Watson et al. (1980). Numbers of species for each genus are from Bryk (1930). 


Genus Number of included spp. 


Leptactea Prout 
Mitradaemon Butler 
Josia Hitibner 

Scea Walker 
Thirmida Walker 
Cyanotricha Prout 
Phavaraea Walker 
Scedros Walker 
Getta Walker 
Anticoreura Prout 
Polyptychia Felder 


Total number of species 103 


— D> 


NrFWOrNN DN OO Or 


Previous systematic studies have shown that Josia belongs in a large, 
well-defined clade that includes other dioptine genera. Below, we re- 
view the evidence supporting that hypothesis. 


The Tribe Josiini 


Most notodontid adults have a relatively simple metathoracic tym- 
panum (Surlykke 1984, Scoble 1992) characterized by a shallow dorsal 
invagination of the epimeron, with the tympanal membrane above, 
facing postero-ventrally. Richards (1932), in his classic study of noctuoid 
tympanal morphology, described a unique type, which he termed the 
“kettle-drum” tympanum, found in the dioptine genera Cyanotricha 
and Josia. In these genera, the epimeron is greatly invaginated to form 
a large, enclosed chamber. The tympanal membrane, oriented hori- 
zontally, forms the ceiling of the chamber (the “head” of the kettle- 
drum). The chamber opens to the outside by means of a small lateral 
hole. The kettle-drum tympanum is figured in Borner (1939: fig. 41), 
Sick (1940: figs. 1-4), Kiriakoff (1950: figs. 9, 10), and Miller (1991: fig. 
240). 

After studying tympana throughout the Dioptinae, Sick (1940) cre- 
ated “Group V”’ for those genera with a kettle-drum tympanum. Group 
V contained eight genera: Mitradaemon Walker, Josia Hiibner, Scea 
Walker, Thirmida Walker, Cyanotricha Prout, Phavaraea Walker, Get- 
ta Walker, and Polyptychia Felder. Kiriakoff (1950) also noted the 
highly derived tympanum of Josia. He divided the “Dioptidae”’ into 
two subfamilies: the Dioptinae with a simple tympanum and the Josiinae 
with a kettle-drum tympanum. Although Kiriakoff examined fewer 


340 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


ons ey lee i 2, 
. 1 . 

\ 1 

. 


) ‘ 
= ° 
1 ae a-ona-"y ce S 
\ ao a a i etaie 
ee : . S.@  «-” Caraca os 
\ Pe i es , : naa 
“ ‘ \ > Maracay we Y *---->- Seger! 
4 Heap Aon Fe —o) =e ‘ 
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\ t SE Ne SC A 
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Recs ge Pres 


Fic. 1. Map of Venezuela; state boundaries shown with dashed lines (scale bar = 100 
km). 


taxa than Sick, his list of genera for the Josiinae corresponds closely 
with Group V. Recent findings have shown that, rather than being a 
separate family as previous authors had supposed, the Dioptinae is a 
highly derived subfamily within the Notodontidae (Minet 1983, Weller 
1989, Miller 1991, 1992b). Kiriakoff’s divisions thus can be recognized 
as tribes, the Dioptini and Josiini (new status). 

We performed an exhaustive survey of tympanal structure in the 
Dioptinae. In addition to the eight genera in Sick’s (1940) Group V, 
we discovered three monobasic genera with the kettle-drum tympanum: 
Leptactea Prout, Scedros Walker, and Anticoreura Prout. According 
to our definition, the tribe Josiini includes 108 described species in 11 
genera (Table 1). 


Previous Works on the Josiini 


Although early authors did not formally recognize the Josiini, they 
nevertheless provided relevant taxonomic treatments of the group. The 


VOLUME 48, NUMBER 4 341 


two most important are Prout’s (1918) preliminary revision of dioptine 
genera and Hering’s (1925) contribution to Seitz, where species diag- 
noses are provided and adults of many taxa are figured. Two keys to 
Josia adults are available, one in Walker (1854: 289-290), and a more 
complete one in Forbes (1981). Forbes (1939) also gave keys and di- 
agnoses for five josiines from Barro Colorado Island, Panama. 

Morphological and biological data are scant. Markin et al. (1989) 
described the biology and life stages of Cyanotricha necyria Felder, a 
josiine species from the western Andes. Miller (1988) described the 
genitalia and copulatory mechanism of C. necyria, and also has dis- 
cussed various features of josiine larval and adult morphology within 
the context of a reclassification of the Notodontidae (Miller 1991). 
Papers by other authors give only anecdotal information on immatures. 
Because of the paucity of published work on the Josiini, we have chosen 
to present considerable detail in this paper. 


METHODS 


During the course of our research we visited sites in mountain systems 
of the Cordillera de la Costa in the state of Aragua of northern Ven- 
ezuela, and in the Cordillera de Mérida in western Venezuela, specif- 
ically in the states of Mérida and Barinas (Fig. 1). Our collecting covered 
a range of elevations (540 to 2420 m), life zones, and vegetation types. 
We use Holdridge’s life zone system, relying on the work of Ewel et 
al. (1976) to identify life zones for each of our collecting sites. Life 
zone types are indicated by an abbreviation in parentheses after the 
locality. The altitudes provided are plus or minus 50 m. 

Most of our collecting sites showed varying degrees of habitat dis- 
turbance, usually due to agriculture. Large tracts of land in the Cor- 
dillera de Mérida and Cordillera de la Costa have a long history of 
agricultural use, and the vegetation currently present, even in some 
forested sites, may be secondary. Primary vegetation is often reduced 
to relictual patches or is limited to steep mountain slopes. Four of our 
collecting sites are within the limits of national parks, Parque Nacional 
Henri Pittier in the state of Aragua, and Sierra Nevada in Merida. In 
these officially protected areas, the vegetation is likely to be primary. 

Passiflora leaves with eggs or larvae of Josiini were picked in the 
field and transported to the laboratory inside moistened plastic con- 
tainers. In the lab, leaves were transferred into small glass flasks con- 
taining water, submersing the petiole, and plugging the neck of the 
flask around the petiole with wet paper. Each flask was placed in a 
plastic dish and covered with an inverted 1 liter transparent plastic 
container. Leaves usually remained fresh in the rearing containers for 


342 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


several days. Fresh food was provided when leaves were consumed or 
showed signs of deterioration. 

For two of the species, Josia radians and J. aurifusa, adult females 
were collected in the field, placed in plastic bags with host foliage, and 
brought to the lab. Females laid eggs on the enclosed leaves over a 
period of two to three days. | 

Larvae were fed leaves of the Passiflora upon which they were 
discovered in the field, or on an alternative species when the original 
host was not easily available. Certain of the josiines lay eggs in batches. 
For these, egg masses were divided and reared in more than one con- 
tainer. Frass accumulations were discarded regularly. Instar durations 
were not monitored precisely; we therefore give only the approximate 
development time from field-collected egg to adult emergence. At least 
for the first, second, and third instars, each stadium lasts approximately 
five or six days. 

We preserved specimens of eggs, first and last instar larvae, and 
adults. Larvae were preserved by dropping them into hot water (just 
below boiling temperature) for several seconds, and then transferring 
them to vials of 70% ethanol. Shed head capsules were saved in alcohol 
along with the associated specimens. Pupal exuviae were mounted on 
the same pin as the adult voucher. Voucher material is deposited in 
the collections of the Museo del Instituto de Zoologia Agricola, Maracay 
(MIZA) and the American Museum of Natural History, New York 
(AMNH). 

Identifications of adult moths were confirmed by comparison with 
type specimens. Comparisons between genitalic dissections of vouchers 
and types were required for Josia radians, J. aurifusa, and J. turgida. 
These three belong to difficult species complexes, the resolution of which 
will require study of large series from a wide range of localities. This 
was beyond the scope of the present work. 

The Passifloraceae contains twenty genera and approximately 600 
species (Heywood 1979). Over 400 species are placed in the genus 
Passiflora, with all but 20 occurring in the Neotropics (Holm-Nielsen 
et al. 1988, Gentry 1998). Stephan S. Tillet (Herbario Ovalles, Facultad 
de Farmacia, Universidad Central de Venezuela, Caracas) verified host- 
plant identifications. We assign Passiflora species to subgenera following 
Killip (1938), who recognized 22 subgenera for the American flora. 
However, the classification of Passiflora currently is undergoing revision 
(J. MacDougal and C. Feuillet, pers. comm.), so these assignments are 
provisional. Plant distributions are based on Killip (1988) and Holm- 
Nielsen et al. (1988). 

Nomenclature for larval morphology follows Stehr (1987) and Pe- 


VOLUME 48, NUMBER 4 343 


terson (1962). We employ the system for larval chaetotaxy developed 
by Hinton (1946), but have incorporated minor modifications suggested 
in Miller (1991) and Rawlins (1992). For first instars we use the setal 
nomenclature of Kitching (1984) on Danaini (Nymphalidae) and Flem- 
ing (1960) on Heliconiinae. Pupal terminology follows Mosher (1916). 
Larval measurements were made using a Microcode II (Boeckeler In- 
struments) digital micrometer attached to a Zeiss SV8 stereomicroscope. 
Electron micrographs were taken with a Zeiss DSM 950 Digital Scanning 
Microscope. 

Abbreviations. A = abdominal segment; CuA = cubital wing vein; 
D = dorsal seta; DC = discal cell; FW = forewing; HW = hindwing; 
JSM = James S. Miller; L = lateral seta, LDO = L. Daniel Otero; LM- 
mf = lower montane moist forest; LM-wf = lower montane wet forest; 
M = medial wing vein; P = posterior seta; PM-df = premontane dry 
forest, PM-mf = premontane moist forest, PM-wf = premontane wet 
forest; R = radial wing vein; SD = subdorsal seta; SV = subventral seta; 
Tl = prothoracic segment; T2 = mesothoracic segment; T3 = meta- 
thoracic segment; T-mf = tropical moist forest; TS = tarsal seta; V = 
ventral seta. (Additional abbreviations are given in the figure legends.) 

Acronyms for Museums. AMNH = American Museum of Natural 
History, New York, NY; BMNH = The Natural History Museum, Lon- 
don, Great Britain, CMNH = Carnegie Museum of Natural History, 
Pittsburgh, PA; CUIC = Cornell University Insect Collections, Ithaca, 
NY; LACM = Los Angeles County Museum of Natural History, CA; 
MIZA = Museo del Instituto de Zoologia Agricola, Maracay, Venezuela; 
MNHN = Muséum National d’Histoire Naturelle, Paris, France; NUNH 
= National Museum of Natural History, Washington, DC; ZMH = 
Zoologisches Museum, Humboldt-Universitat, Berlin, Germany. 


GENERAL ACCOUNT 


Many of the traits we observed are found throughout our study taxa. 
General morphological features, as well as those that provide potentially 
useful taxonomic differences, are summarized below. 


Egg. The pearly white eggs are almost perfectly spherical (Figs. 2, 15), without surface 
sculpturing except for a fine reticulate pattern in the area surrounding the micropyle 
(Figs. 3, 4). Among the five study species, egg size ranges from 0.70 to 1.10 mm in 
diameter. They are laid either individually, or in small clusters of up to 25 eggs (Fig. 
15). 

First Instar Larva. All first instars in this study exhibit the same simple primary setal 
pattern (Figs. 47, 56, 71, 78); setae L2 and L8 absent on segments T2 and T8, L8 absent 
on Al-—A7. This configuration also occurs in first instars of Nymphalidae (Kitching 1984), 
Heliconiinae (Fleming 1960), and Noctuidae (Merzheevskaya 1988), and is probably 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 2-7. Scanning electron micrographs of Thirmida discinota immature stages. 2, 
Egg, dorsal view (x79); 3, Micropyle and surrounding area (x 1268); 4, Micropyle ( x6960); 
5, Head and T1 of first instar, lateral view (x97); 6, Tarsus of first instar T3 leg, mesal 
view (596); 7, A8 spiracle and setal bases, first instar (x 344). [Pg = prothoracic gland.] 


VOLUME 48, NUMBER 4 345 


distributed throughout the Lepidoptera. Features restricted to first instar larvae include: 
antenna short (Figs. 5, 48, 61, 79); spinneret long and apically acute (Figs. 5, 48, 61, 79); 
mandible serrate (Godfrey et al. 1989); tarsal setae elongate (Figs. 6, 49); each primary 
seta located on a prominent, sclerotized pinaculum (Figs. 5, 7, 47, 56, 71, 78); spiracles 
extremely small, round (Figs. 5, 7); color pattern simple, either entirely white or with 
alternating reddish bands (compare Figs. 47 and 56). 

Fourth (Final) Instar Larva. Subprimary setae are defined as those that appear after 
the first larval molt (Hinton 1946, Kitching 1984). In the josiines we studied, second, 
third, and fourth instars have the same setal pattern. However, patterns can vary between 
species, suggesting that subprimary setae may be useful in josiine systematics. For example, 
fourth instars of Josia gopala possess an L seta on segments A2—A6 not found in other 
species (compare Figs. 52 and 67). In Thirmida discinota, L3 on A3-A6 is multiple, and 
there are novel setae on the anal plate (Figs. 48, 44). Characters such as these may provide 
useful synapomorphies for defining subclades within the Josiini. Color pattern differences 
provide important characters for separating species. 

A diagnosis of last instar larval morphology for Josiini is as follows: Head either entirely 
black (Fig. 45), or white with dark dorso-ventral stripes (Figs. 54, 69, 76, 85); mandibular 
margin smooth (Godfrey et al. 1989); antenna with segment 2 longer than segment 1, 
segment 3 short (Figs. 10, 51, 80, 81); spinneret short and wide (Figs. 11, 50, 62). Primary 
setal pattern generally as in other Notodontidae; secondary setae occasionally present on 
head and body (in Cyanotricha necyria; Miller 1991); tarsal setae TS2 and TS4 lanceolate 
with apices acute, TS3 broad, leaf-shaped (Figs. 12, 18, 57, 63, 82); prothoracic gland 
present (Figs. 8, 9, 43, 52, 67, 74, 83); thorax and abdomen lacking projections, but with 
a slight hump on A8 (Figs. 44, 53, 68, 75, 84); cuticle glossy in appearance, covered with 
microprojections (= shagreened;” e.g., Figs. 5, 7, 20-23); body variously patterned with 
reddish maroon, yellow and white (Figs. 14-19); A8 spiracle either subcircular or an 
elongate oval (Figs. 21-23); terminal segments often conspicuous, white (Figs. 16-19); 
prolegs on A10 reduced but functional (Figs. 44, 53, 68, 75, 84); segments A7-A10 
frequently held aloft (Figs. 16, 18, 19); like other Dioptinae (but unlike other Notodon- 
tidae), a single MD seta on Al rather than two (Miller 1991). 

Pupa. Most features of josiine pupal morphology do not show significant variation (see 
Miller 1992b: figs. 31, 32). However, the cremaster varies in shape, and in the number 
and size of the cremaster setae (Figs. 24, 25, 58-60, 64-66). 

Adult. Many external adult traits show variation, and these can be used in identification. 
Examples include the shape of the labial palpus and length of the pectinations on the 
male antenna. The forewing can exhibit a longitudinal stripe (Figs. 28-36), a transverse 
one (Hering 1925), or variations on the latter (Figs. 26, 27). The species of Josiini also 
differ greatly in size; FW lengths range from 11 mm (Josia ena Boisduval) to 28 mm 
(Phavaraea rejecta Hibner). 


SPECIES ACCOUNTS 
Thirmida discinota Warren, 1900, revised status 


Distribution. This species is known almost exclusively from localities 
close to the city of Mérida. However, a single specimen (MIZA) from 
the State of Lara has the following data: Parque Nacional Yacambt, 
El! Blanquito, 1350 m; 1-3/VIII/1976; C. J. Rosales, L. J. Joly. This is 
the lowest elevation from which T. discinota is known; all the other 
specimens were collected at or above 2000 m. Data for the female 
holotype (BMNH) is given as Pedregosa, Mérida, 3000 m, October 1897 
(Briceno) (Warren 1900:129), a locality southwest of the city of Mérida. 
Thirmida discinota is rare in collections. Previously it was represented 


346 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 8-13. Scanning electron micrographs of fourth instar Thirmida discinota larva. 
8, Head and T1 (x22); 9, Prothoracic gland, lateral view (x 133); 10, Antenna, frontal 
view (180); 11, Maxillary and hypopharyngeal complexes, frontal view (<133); 12, 
Tibia and tarsus of right T2 leg, mesal view (x82); 13, Tarsus of right T2 leg, mesal 
view (x290). [An = antennal segment; Cl = tarsal claw; Pg = prothoracic gland; Sp = 
spinneret; 2, 3, 4 = tarsal setae 2, 3, and 4.] 


VOLUME 48, NUMBER 4 347 


Fics. 14-19. Immature stages of Josiini. 14, Fourth instar larva of Thirmida discinota 
on Passiflora bauhinifolia; 15, Eggs of Thirmida discinota laid on the underside of P. 
bauhinifolia leaves; 16, Fourth instar of Josia radians on Passiflora capsularis; 17, Fourth 
instar larva of Josia gopala on Passiflora cuneata; 18, Fourth instar larva of Josia aurifusa 


on P. capsularis; 19, Fourth instar larva of Josia turgida on P. capsularis. Photographs 
by L. Daniel Otero. 


in major museums by a total of 18 specimens (BMNH, MIZA, NMNH, 
and ZMH), most of which were collected near the turn of the century. 
We have found it to be locally abundant. 

Collecting Localities. Recorded from Quebrada La Cafia on the road 
from El Valle to La Culata, northwest of Mérida, at 2420 m ($1 October 


348 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 20-25. Scanning electron micrographs of Josiini immature stages. 20, A4 spi- 
racle, fourth instar Thirmida discinota larva (note seta MSD2 at left) (x211); 21, A8 
spiracle, fourth instar T. discinota larva (x151); 22, A8 spiracle, fourth instar Josia 
aurifusa larva (x248); 23, A8 spiracle, fourth instar Josia radians larva (243); 24, 
Cremaster of T. discinota pupa, postero-ventral view (x58); 25, Cremaster of T. discinota, 
dorsal view (x59). 


VOLUME 48, NUMBER 4 349 


1992, 17 January 1993, and 6 April 1993). The life zone for this locality 
is regarded as lower montane moist forest (LM-mf) according to Ewel 
et al. (1976). However, only relictual patches of the original forest type 
remain, and the habitat where we found T. discinota is characterized 
by secondary vegetation that includes shrubs and some small trees (Fig. 
42). 

Foodplant. Eggs of T. discinota were collected on vines of Passiflora 
bauhinifolia Kunth (subgenus Plectostemma), growing near a stream. 
There is some confusion regarding the distribution and taxonomic status 
of this species; Holm-Nielsen et al. (1988) regard it as a synonym of P. 
alnifolia Kunth, and according to Killip (1938) the plant occurs only 
in southwestern Colombia, Ecuador, and northern Peru. 


Egg. Relatively large (1.1 mm; n = 5); deposited in clusters of 4 to 25 eggs (Fig. 15). 
Eggs tend to be laid on older leaves than in the other Josiini that we observed. Markin 
et al. (1989) noted a similar preference for mature foliage by ovipositing females of 
Cyanotricha necyria Felder (Josiini). Based on field collections, duration of the egg stage 
is at least 10 days. Development time from field-collected egg to adult = 60-65 days. 

First Instar. Head entirely black except for white clypeus; thorax and abdomen un- 
pigmented. Primary setae (Fig. 47) as in general account (above), similar to Josia. Body 
length = 5.24-5.58 mm; head width = 0.77-0.78 mm (n = 5). 

Second and Third Instars. Similar in setal and pigment patterns to final instar larvae. 
Head widths: 2nd instar = 1.14—1.22 mm; 3rd instar = 1.80-1.88 mm (n = 5). 

Final (Fourth) Instar. Head entirely shiny black except for white clypeus and thin 
white lines along ecdysial suture and ecdysial lines (Fig. 45); labrum with a whitish central 
area; antenna with segment 2 expanded distally, approximately twice the length of 
segment 1. Prothoracic shield (Fig. 46) broad, heavily sclerotized, with a partial medial 
seam, anterolateral angles not produced; ground color of thorax and abdomen reddish 
purple (Fig. 14); a lemon yellow longitudinal stripe along dorsal midline bordered on 
each side by a thin, irregular whitish subdorsal stripe; lateral pattern complex (Figs. 43, 
44), composed of a wide yellow longitudinal stripe with reddish purple blotches and an 
irregular reddish purple stripe within it; venter entirely lemon yellow; segment AQ not 
conspicuous, mostly maroon above. Tarsal setae with TS2 and TS4 lanceolate, TS3 broad 
and leaf-shaped, apex emarginate (Fig. 13). Setal pattern (Figs. 43, 44) on thorax and 
abdomen similar to other Josiini, but with the following exceptions: seta SV2 on segment 
A2 located between L3 and SV3, on a horizontal line with them; one or two additional 
setae present on proleg-bearing segments (A3—A6) in the L3 position; anal plate with 2- 
8 additional setae on each side; lateral plate of Al0 proleg base with more than 12 setae; 
all primary setae wide, white and “fleshy” in appearance; each thoracic and abdominal 
seta on a small, heavily sclerotized pinaculum. Body length = 38.0-39.0 mm; head width 
= 2.60-2.83 mm (n = 5). 

Pupa. Cremaster (Figs. 24, 25) short and broad, heavily rugose dorsally with broad, 
longitudinal flutes ventrally; approximately 20 short, hook-shaped setae. Duration: ap- 
proximately 20 days. 

Adult. FW length = 23-25 mm. Head, thorax and abdomen gray-brown, abdomen 
with scattered iridescent bluish scales; pectinations of male antenna long; eye relatively 
small; front and labial palpus rough-scaled; palpus porrect, narrow, fringed below with 
long scales; pectus and femur with long, hairlike scales. FW light orange from base to 
distal end of DC, charcoal gray beyond (Figs. 26, 27); orange region with diffuse, black 
longitudinal lines along veins, surrounded on anterior, posterior, and distal margins by a 
black border; HW (Figs. 26, 27) with a light orange central region extending from base 
to slightly beyond DC, iridescent purple-black merging to charcoal gray beyond; cubital 


390 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


27 
Sd. 
3 
3 


1 


OF 


36 


bd 
>. 
BE 
=? 


Fics. 26-36. Adults of Josiini (life size). 26, Thirmida discinota, male; 27, T. dis- 
cinota, female; 28, Josia radians, male; 29, J. radians, female; 30, Josia gopala, male; 
31, J. gopala, female; 32, Josia turgida, male; 33, J. turgida, female; 34, Josia aurifusa, 
male (dark form from Puente Victoria); 35, J. aurifusa, male; 36, J. aurifusa, female. 
Photograph by Cal Snyder. 


vein thinly lined with black scales; a small, black spot on upper discocellular cross vein. 
Upper and lower wing surfaces the same, except veins below uniformly orange rather 


than black. 


Discussion. Warren (1900) described discinota in the genus Scea, 
which was at that time placed in the Cyllopodinae (Geometridae). His 
description was based on a single female specimen (BMNH). Warren’s 
taxon was transferred to Thirmida in the Dioptinae by Prout (1918), 


VOLUME 48, NUMBER 4 Sol 


Fics. 37-42. Collecting sites for Josiini (all in the State of Mérida, Venezuela). 37, 
Paseo Los Pinos (1450 m), a locality for Josia radians; 38, Along the road from Estanques 
to Paramo Las Coloradas (1150 m), a locality for J. radians; 39, Las Playitas, near 
Bailadores (2250 m), a locality for J. radians (the foliage and flowers at left are those of 
Passiflora manicata); 40, Cloud forest margin (2300 m) in Monterrey, a locality for J. 
gopala; 41, Puente Victoria (540 m), a locality for J. aurifusa; 42, La Cana on the road 
from El Valle to La Culata (2420 m), northwest of the city of Mérida, a locality for 
Thirmida discinota. Photographs by L. Daniel Otero. 


who considered discinota to be a subspecies of T. dimidiata Walker. 
This arrangement was followed by all subsequent authors (Hering 1925, 
Talbot 1929, Bryk 1930). We compared specimens of T. dimidiata 
from Bogota, Colombia, the type locality (Walker 1854:466), with our 
Merida material, and discovered numerous differences in wing pattern 
and genitalia. We therefore recognize the two as distinct, hereby re- 
turning Thirmida discinota to species level status. 

Thus we recognize six species in Thirmida, five of which are figured 
in Hering (1925: plate 71). The only one with which T. discinota could 
be confused is T. dimidiata. Adults may be separated on the basis of 


302 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 48-47. Larval stages of Thirmida discinota. 43, Head, thorax, and segments 
A1-A3 of fourth instar, lateral view; 44, Segments A6-A10 of fourth instar, lateral view 
(scale line = 2 mm); 45, Head of fourth instar, frontal view (scale line = 1 mm); 46, 
Prothoracic shield of fourth instar, dorsal view (scale line = 1 mm); 4.7, First instar, lateral 
view (scale line = 1 mm). [See Figs. 52-56 for key to symbols. ] 


VOLUME 48, NUMBER 4 | 358 


wing pattern: the orange in the forewing of T. dimidiata extends beyond 
the distal margin of the discal cell, while in T. discinota it stops at the 
end of the discal cell (Figs. 26, 27). 

This life history description is the first for a member of Thirmida. 
Adults and immatures of the other species, known from the Andes of 
Colombia and Ecuador at elevations between 1200 and 2300 m, will 
likely be discovered by collecting in habitats similar to that of T. dis- 
cinota. Hering (1925), noting wing pattern similarities, suggested that 
the closest relatives of Thirmida are the Andean genera Scea and Cy- 
anotricha. 

Mimicry occurs throughout the Dioptinae (Seitz 1925, Kohler 1930). 
In his description of the new genus and species Thermidarctia ther- 
midoides in the Pericopinae (Arctiidae), Talbot (1929) noted what he 
called an “almost perfect resemblance” (p. 183) between his taxon and 
Thirmida discinota. Mérida is the type locality for both moths. 


Josia radians Warren, 1905a 


Distribution. Bryk (1930) reported J. radians from Mexico to Co- 
lombia, Guyana, and Venezuela, presumably based on museum spec- 
imens. However, J. radians frequently is confused with J. ligata Walker 
(type locality Bogota, Colombia) and J. frigida Druce (type locality 
Las Mercedes, Guatemala). In all probability, the majority of material 
that Bryk examined was misidentified. 

Reliably determined museum material, and specimens we have col- 
lected, suggest that the species occurs from the State of Aragua, Ven- 
ezuela west to the Colombian border. It probably occurs in eastern 
Colombia as well. Although there is no locality data on the holotype, 
Warren’s (1905a) description of J. radians gave the following: “1 male 
from Onaca, Sta. Martha, 2200 ft., wet season, September-October 1901 
(Engelke).” We have been unable to locate this site, but assume that it 
is a misspelling of Ocania, a locality in the eastern Cordillera of Colombia 
near the Venezuelan border (Fig. 1). 

Collecting Localities. Eggs, larvae, and adults of Josia radians were 
collected at eight different localities (Figs. 37-39, 72). There were seven 
sites in the state of Mérida, and an eighth locality in the Parque Nacional 
Henri Pittier, State of Aragua. Elevations ranged from 1100 m (near 
El Amparo) to 2250 m (Las Playitas, near Bailadores). These collecting 
sites cover a variety of life zones (PM-mf, PM-wf, PM-df) and vegetation 
types. They include cloud forest (Henri Pittier), as well as highly dis- 
turbed habitats such as a coffee plantation (La Mucuy) and a small 
patch of Passiflora on the road margin in agricultural land (Las Playitas). 
In one case (Paseo los Pinos near the city of Mérida), J. radians im- 


304 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


matures were found on their host in a vacant lot within a residential 
area. 

The species seems to occur throughout the year; our collecting dates 
include the months of May, June, July, September, October, and De- 
cember of 1992, as well as February and March of 1993. 

Foodplant. We collected Josia radians on four Passiflora species. It 
was most frequently found on P. capsularis L. (subgenus Plectostem- 
ma), a widespread species that occurs from Guatemala south to central 
Brazil and Paraguay up to 1900 m, and throughout the Greater Antilles. 
At Las Playitas (Fig. 39), the highest elevation so far known for J. 
radians (2250 m), larvae were collected on Passiflora manicata Jussieu 
(subgenus Granadillastrum). This plant is known from western Ven- 
ezuela east to Colombia and south to northern Peru between 1500 and 
2500 m elevation. The third Passiflora used by J. radians is P. cuneata 
(subgenus Plectostemma), one of the hosts for Josia gopala (see below). 
Eggs of J. radians also were found on P. rubra L. (subgenus Plectos- 
temma). Passiflora rubra is widespread, occurring throughout the West 
Indies, from Venezuela and Colombia south to Bolivia, and in eastern 
Brazil. Passiflora rubra and P. capsularis are close relatives and difficult 
to distinguish (Killip 19388, Holm-Nielsen 1988, Vanderplank 1991). 
Our identifications were confirmed by comparison of flowers and fruits. 


Egg. Diameter = 0.91-0.98 mm (n = 12). Eggs are laid singly. Duration: 7 to 8 days. 
Development time from egg to adult = 40-45 days. 

First instar. Head entirely black, except clypeus white. First instars of J. radians differ 
from the other species we studied in the following: antennal segments 1 and 2 short (Fig. 
48); thoracic legs with tarsal setae elongate, apex of TS3 emarginate (Fig. 49); thorax 
with a red dorsal patch on T2 (Fig. 56); segments Al, A3 and A8 ringed with reddish, 
segments A5 and A10 with a red dorsal patch, A7 with a red lateral patch below spiracle 
(Fig. 56). Body length = 5.73-6.01 mm (n = 3); head width = 0.54-0.57 mm (n = 6). 

Second and Third Instars. Similar in color and setal pattern to final instar. Head widths: 
2nd = 0.90-0.91 mm (n = 6); 38rd = 1.40-1.44 mm (n = 6). 

Final (Fourth) Instar. Head shiny black with white front, wide white regions extending 
along ecdysial lines down to antenna, and a pair of lateral white stripes, each from vertex 
to stemmata, widening below (Fig. 54); antenna with segment 2 approximately twice the 
length of segment 1 (Fig. 51). Prothoracic shield with anterolateral angles produced, an 
unpigmented medial seam (Fig. 55); ground color of thorax and abdomen reddish maroon, 
with a complex white and yellow pattern overlying it (Fig. 16); lateral and dorsal light- 
colored areas reticulate (Figs. 52, 53); venter entirely white; dorsal red portions more 
heavily pigmented in an alternating pattern on segments T2, Al, A3 and A5; segment 
A9 conspicuous, entirely white above (Figs. 16, 53). Tarsal setae (Fig. 57) similar to 
Thirmida discinota. Pattern of primary setae on thorax and abdomen (Figs. 52, 53) similar 
to J. aurifusa and J. turgida; lateral plate of Al0 proleg base with approximately 9 setae. 
Body length = 28-30 mm; head width = 2.18-2.22 mm (n = 5). 

Pupa. Cremaster (Figs. 58-60) conical, with a row of longitudinal striae circling base 
and two more distal concentric rings of reticulate striae; eight stout, hook-shaped setae. 
The pupal stage lasted 14 days. 

Adult. FW length = 16.0-19.0 mm. Head orange-yellow with vertex, antenna and 
scales behind eye black; pectinations of male antenna moderately long; labial palpus 


VOLUME 48, NUMBER 4 355 


Fics. 48-51. Scanning electron micrographs of Josia radians larvae. 48, Ventral 
portion of first instar head, lateral view (<254); 49, Tarsus of right metathoracic leg of 
first instar, mesal view, showing tarsal setae (x 800); 50, Ventral portion of fourth instar 
head, frontal view of right side (x68); 51, Right antenna of fourth instar, frontal view 
(x 200). [An = antenna; Cl = tarsal claw; Lb = labrum; Mx = maxillary palpus; Sp = 
spinneret; 2, 3, 4 = tarsal setae 2, 3, and 4.] 


porrect, exceeding front; palpus relatively wide, segment 1 yellow, segment 2 black with 
yellow scales on venter, segment 3 black. Dorsum of thorax black with orange-yellow 
stripe on each side running from patagium to abdomen, tegula black in lateral half, yellow 
along mesal half, with fringe of long yellow scales distally; lateral portions of thorax 
mostly black, orange-yellow below wing bases; legs black, buff scales on dorsal surfaces; 
anterior surfaces of forecoxa light buff. FW and HW (Figs. 28, 29) black with thin, 
longitudinal orange-yellow stripe from base to within 1 mm of outer margin, stripe slightly 
wider in female; FW and HW costa orange-yellow from base to approximately two thirds 
out; HW anal margin orange-yellow; longitudinal stripe of FW wider and more diffuse 
below. Abdomen black with orange-yellow lateral stripe on each side, stripe becoming 
gray at caudal margin of A8; venter of abdomen whitish buff. 


Discussion. The moths we reared match the male holotype of Josia 
radians Warren (BMNH) in wing pattern and genitalic morphology. 


356 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


sp1 / / Se 


Pree 
me 


L3 


Fics. 52-56. Larval stages of Josia radians. 52, Head, thorax, and segments Al—-A8 
of fourth instar, lateral view; 53, Segments A6-A10 of fourth instar, lateral view (scale 
line = 2 mm); 54, Head of fourth instar, frontal view (scale line = 1 mm); 55, Prothoracic 
shield of fourth instar, dorsal view (scale line = 0.5 mm); 56, First instar, lateral view 
(scale line = 1 mm). [A = anterior seta; Af = adfrontal seta; C = clypeal seta; E = epicranial 
suture; Ed = ecdysial line; F = frontal seta, MD = dorsal proprioceptor seta; P = pos- 
teriodorsal seta; Pg = prothoracic gland; S = stemmatal seta; Sp = spiracle; XD = XD 
seta; for other symbols see “Methods. ”’] 


VOLUME 48, NUMBER 4 Sloe 


Fics. 57-60. Scanning electron micrographs of immature stages of Josia radians. 57, 
Tarsus of right T3 leg of fourth instar larva, mesal view, showing tarsal setae 2, 3, and 4 
(<358); 58, Cremaster, posterior view (x69); 59, Cremaster, dorsal view (x73); 60, 
Cremaster, ventral view (x71). [Cl = tarsal claw. ] 


However, J. radians belongs to a group of closely related species that 
includes J. ligata Walker, J. frigida Druce, and J. fustula Warren. 
Identifications in this complex are extremely difficult (Forbes 1931). 


Josia gopala Dognin, 1891 


Distribution. Josia gopala is known only from the State of Merida, 
Venezuela (AMNH, BMNH, MIZA, NMNH, ZMH). The locality label 
on the female holotype (NMNH) reads “Mérida, Venezuela; Terre 
Temperee,” the latter (in French) probably referring to a temperate 
habitat. Based on museum label data and on data we have accumulated, 
Josia gopala occurs within a fairly restricted altitudinal range, between 
2000 and 3000 meters. Although J. gopala is rare in museum collections, 


358 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


previously known from less than 20 specimens, the moth can be quite 
common in Merida. 

Collecting Localities. Eggs and larvae were collected at four cloud 
forest localities. Two sites, close together in Asentamiento Monterrey, 
El Valle (LM-mf) at 2300 and 2350 m elevation, were visited on 14 
and 23 June 1992, 16 August 1992, and 7 March 1993. One of these is 
along a cloud forest margin bounded by agricultural land and dispersed 
human habitations (Fig. 40), while the other is in a pine plantation in 
what was formerly cloud forest. Our other two sites were along the 
road to El Morro (2150 m), visited on 6 September 1992 (LM-mf), and 
in Parque Nacional Sierra Nevada, La Mucuy (2300 m), visited on 12 
October 1992 (LM-wf). The species seems to be restricted to the borders 
of forested habitats. 

Foodplant. Josia gopala was discovered feeding on two Passiflora 
species. Eggs were collected on Passiflora cuneata Willdenow (subgenus 
Plectostemma) growing in the shaded forest margins. This Passiflora 
is known from the mountains of central Venezuela east to the Cordillera 
Santa Marta and eastern Cordillera of Colombia at elevations between 
700 and 3800 m. Caterpillars also were found on Passiflora gritensis 
Karsten (subgenus Granadilla). In one case, the P. gritensis plants were 
growing in a pine plantation. Passiflora gritensis is known only from 
western Venezuela at altitudes near 2500 m. These two Passiflora spe- 
cies occur together at the Monterrey sites. Females of J. gopala tend 
to oviposit on tender, lower leaves of the host near the ground, especially 
on the leaves of seedlings or new shoots. 


Egg. Diameter = 0.86-1.01 mm (n = 18). Duration: 7 to 8 days. Eggs are laid indi- 
vidually, not in clusters. Development time from egg to adult = 52-56 days. 

First Instar. Head dark brown except for white clypeus, and small light brown patches 
at ventral angles of front beyond ecdysial lines. General morphology and primary setae 
(Figs. 61, 71) similar to other Josiini. Body length = 4.388-5.19 mm; head width = 0.57- 
0.62 mm. 

Second and Third Instars. Similar in pigmentation and setal pattern to final instar. 
Head widths: 2nd = 0.89-0.95 mm; 8rd = 1.39-1.50 mm (n = 4). 

Final (Fourth) Instar. Head shiny black except for white frons and clypeus, and two 
ovoid white patches on either side of ecdysial lines (Fig. 69); antenna with segment 2 
approximately twice the length of segment 1. Prothoracic shield narrow, no medial seam 
(Fig. 70); ground color of thorax and abdomen reddish maroon, overlying pattern simple, 
white and yellow (Fig. 17); light-colored lateral and dorsal areas roughly rectangular in 
shape (Figs. 67, 68); venter entirely white; segment A9 conspicuous, entirely white above. 
Tarsal setae (Fig. 63) with TS2 lanceolate, TS3 broad and leaf-shaped, TS4 sword-shaped 
with apex acute. Setal pattern on T1—A10 (Figs. 67, 68) similar to last instars of other 
Josia species except: a novel L seta (seta “Lx” in Fig. 67) present on segments A2—A6, 
located between SD1 and L1]; lateral plate of Al0 proleg base with only 4 setae (n = 9 
in other Josia species). Body length = 25-26 mm; head width 1.9-2.13 mm (n = 5). 

Pupa. Cremaster (Figs. 64-66) short, somewhat flattened distally, fluting irregular; 
eight delicate, widely spaced, hook-shaped setae. Duration: approximately 19 days. The 
pre-pupal larvae weave a shelter of leaves and crawl into it to pupate (2 observations). 

Adult. FW length = 16.5-17.5 mm. Head, thorax, and abdomen dark charcoal blue- 


VOLUME 48, NUMBER 4 359 


Fics. 61-66. Scanning electron micrographs of immature stages of Josia gopala. 61, 
Ventral portion of first instar head, lateral view ( x 294); 62, Maxillary and hypopharyngeal 
complexes of fourth instar larva, frontal view (215); 63, Tarsus of right prothoracic leg 
of fourth instar larva, mesal view (468); 64, Cremaster, posterior view (x81); 65, 
Cremaster, dorso-posterior view (x76); 66, Cremaster, ventral view (x81). [Cl = tarsal 
claw; Hp = hypopharynx; Sp = spinneret. ] 


360 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


D1 Di D2 
Soe A 
7 >! ‘ ae \ eee ft re ee {soi [ ee lls ee ge 


? 


Fics. 67-71. Larval stages of Josia gopala. 67, Head, thorax, and segments Al—-A38 
of fourth instar, lateral view; 68, Segments A6—A10 of fourth instar, lateral view (scale 
line = 2 mm); 69, Head of fourth instar, frontal view (scale line = 1 mm); 70, Prothoracic 
shield of fourth instar, dorsal view (scale line = 0.5 mm); 71, First instar, lateral view 
(scale line = 1 mm). [Lx = novel L seta; see text. ] 


gray, a lighter lateral stripe on each side of abdomen; pectinations of male antenna short; 
labial palpus narrow and porrect, slightly exceeding front. Wings (Figs. 30, 31) with outer 
margins dark bluish black. FW with a wide, yellow longitudinal stripe extending from 
base to fork of M, and CuA1, just beyond DC; posterior margin light gray below; costa 
black; HW with large yellow central area; anterior margin light gray above; dorsal and 
ventral wing surfaces with essentially the same pattern. 


VOLUME 48, NUMBER 4 | 361 


Discussion. The species gopala was described by Dognin (1891) in 
Flavinia, a genus in the Geometridae (Watson et al. 1980), but was 
moved by Prout (1918) to Josia. The moth can be separated from most 
other Josia by the wide, truncate, longitudinal FW stripe which extends 
for only two thirds the wing length (Figs. 30, 31; Hering 1925). The 
larva exhibits a relatively simple pattern, with roughly rectangular, 
yellow and white patches on each segment laterally and dorsally (Fig. 
17). The last instar is unique among the known species of Josiini in its 
possession of an extra L seta on segments A2-A6 (Figs. 67, 68). 

Based on similarities of wing pattern (Forbes 1931) and genitalia 
(JSM, unpubl.), the closest relatives of Josia gopala are J. patula Walker 
and J. gephyra Hering. These two are slightly larger species (FW length 
= 18.5-21.0 mm) with the yellow forewing stripe narrower and ex- 
tending further out (Hering 1925). Josia patula is fairly well represented 
in museum collections, being known from Choachi, Villavicencio, and 
Bogota, Colombia (BMNH, CMNH, CUIC, NMNH, ZMH). Josia ge- 
phyra is rare; we have seen only four specimens (NMNH, BMNH), all 
collected at the type locality, Cafion del Monte Tolima, Colombia. This 
is a high altitude site; one of these specimens was caught at 4700 m 
(BMNH). Assuming the label is correct, this is the highest altitude 
recorded for any species of Dioptinae. 


Josia aurifusa Walker, 1854 


Distribution. Josia aurifusa appears to be endemic to Venezuela, 
occurring from the state of Aragua west to Tachira (AMNH, CMNH, 
MIZA, MNHN). The label on the male holotype (BMNH) gives the 
locality simply as “Venezuela.” 

Collecting Localities. We collected eggs, larvae, and adults of Josia 
aurifusa at two localities in the Parque Nacional Henri Pittier (PM- 
wf) in the State of Aragua. On 22 March 1992, we collected six eggs 
on Passiflora leaves at Rancho Grande Field Station (1100 m). We also 
collected 16 eggs and one final instar larva on 2 April 1992, near a 
stream crossing the road from Maracay to Choroni (860 m) on the 
northern slope of the mountains (Fig. 73). An adult female, collected 
at the second locality, produced eight eggs from which reared material 
was obtained. Eggs of the species also were collected at a third locality 
on the road from Barinas to Santo Domingo at Quebrada El Alambique 
(1250 m; also PM-wf) in the State of Barinas. All three are cloud forest 
sites. Eggs and larvae of what turned out to be an extremely dark form 
of Josia aurifusa (Fig. 34) were found in the state of Mérida at Puente 
Victoria (540 m, T-mf) on the road from Mérida to El Vigia (Fig. 41). 


362 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Foodplant. Josia aurifusa was found on two Passiflora species. At 
the Aragua localities it fed on P. rubra, while at Puente Victoria it was 
on P. capsularis. Both plant species are discussed under J. radians 
(above). 


Egg. Diameter = approximately 0.7-0.8 mm; laid either singly or in small clusters of 
up to nine eggs. Duration: 4-5 days. Development time from egg to adult = 59 days. 

First Instar. Head dark brown, slightly lighter on front, antenna short (Fig. 79); body 
lacking pigment (Fig. 78). Body length = 4.11 mm (n = 1); head width = 0.49-0.51 mm 
(n = 5). 

Second and Third Instars. Similar in markings and setal pattern to final instar. Head 
widths: 2nd instar = 0.82-0.83 mm; 3rd instar = 1.25-1.80 mm (n = 5). 

Final (Fourth) Instar. Head white, with wide, brownish black frontolateral stripes 
extending from stemmata up to epicranial suture, stripes widening near suture (Fig. 76); 
labrum black with a white central region; antenna (Figs. 80, 81) with segment 2 relatively 
short, less than twice the length of segment 1. Prothoracic shield (Fig. 77) with an 
unpigmented medial seam, anterolateral angles produced, curled inward; ground color 
of thorax and abdomen reddish maroon (Fig. 18), red colored regions lightly spotted with 
white; yellow-white dorsal and lateral patches irregularly shaped (Figs. 74, 75), roughly 
rectangular, relatively small (compared to J. turgida, below); seta L8 on A3-A6 sur- 
rounded by a smallish white spot (Fig. 75); venter white with red, transverse connecting 
lines on Al and A2; segment AQ conspicuous, entirely white above (Fig. 18). Tarsal setae 
as in Josia gopala. Primary setae (Figs. 74-76) as in Josia radians; lateral plate of Al0 
proleg base with approximately 9 setae. Body length = 24.5-26.0 mm; head width = 
1.86-1.98 mm (n = 6). 

Pupa. Cremaster blunt, with uneven longitudinal flutes at base and irregular striae on 
distal flattened portion; strongly concave ventrally; eight hook-shaped setae present. Du- 
ration: 10 days. 

Adult. FW length = 13.5-17.0 mm. Antenna black; head mostly black, face white with 
front blackish gray; scales at antennal base white; pectinations of male antenna moderately 
long; labial palpus porrect, relatively long and thin, extending well beyond front; basal 
three fourths of palpus segment 1 white, rest of palpus black. Dorsum of thorax black, 
an orange stripe on each side running from patagium to base of abdomen; tegula orange, 
black at base; lateral portions of thorax orange; coxa black laterally, white on anterior 
surfaces; legs dark gray, tibiae whitish-buff below. FW black with a wide, orange lon- 
gitudinal stripe from base to within 2 mm of outer margin (Figs. 35, 36), stripe tapering 
gradually toward apex; Rs entirely black through orange stripe; HW (Figs. 35, 36) black 
with caudal two thirds orange; orange region with a diffuse, black, wedge-shaped streak 
along vein 2A from near base, widening toward outer margin; costa of HW light orange 
in basal two thirds. Abdomen black with wide, orange lateral stripes, stripes becoming 
buff at caudal margin of A8; venter white. HW almost entirely black in some specimens 
(Fig. 34), with a diffuse orange streak along CuA2, and anal margin a combination of 
orange and black scales. 


Discussion. Walker’s description of aurifusa (1854: 298) and the 
holotype (BMNH) match our material from Choroni. However, Josia 
aurifusa is a variable species (Hering 1925); our specimens from Puente 
Victoria are extremely dark (Fig. 34). Four names have been synony- 
mized with aurifusa, all of which currently stand as either forms or 
aberrations (Bryk 1930). The phenotypes associated with these names 
vary in the shape of the longitudinal forewing stripe and in the amount 
of black in the hind wing. We recognize one of those, “form” turgida 
Warren, as a distinct species (see below). 


VOLUME 48, NUMBER 4 . 363 


Fics. 72, 73. Collecting sites for Josia species. 72, Beside the road to La Mesa de 
Ejido, State of Mérida, 1.8 km from the turnoff in the Panamerican Highway on the way 
to Jaji (1560 m), a locality for Josia radians; 73, Along the road from Maracay to Choroni 
(860 m), State of Aragua, a locality for Josia aurifusa. Photographs by L. Daniel Otero. 


Josia turgida Warren, 1905b, revised status 


Distribution. Like Josia aurifusa, J. turgida appears to be endemic 
to Venezuela. The species is sympatric with J. aurifusa, but it has been 
recorded from further east (Puerto la Cruz, Anzoatequi; CMNH), and 
does not seem to extend as far west; our western-most record is Altamira 
in the state of Barinas (LDO, collector). The type locality is Valencia, 
Venezuela (State of Carabobo). 

Collecting Localities. Eggs and first instar larvae of Josia turgida 
were found along the road to Altamira in the State of Barinas, ap- 
proximately 2.6 km from the turnoff on the Barinas to Santo Domingo 
road (645 m; T-mf). Eleven eggs were collected on 28 June 1992, and 
14 eggs and five first instar larvae on 13 September 1992. Eggs of the 
species also were collected on the road from Barinas to Santo Domingo 
at Quebrada El Alambique (1250 m) in the State of Barinas. 

Foodplant. The known hostplants are Passiflora capsularis and P. 
rubra. These Passiflora species also are used by J. aurifusa and J. 
radians. Larvae were collected from plants growing in secondary veg- 
etation along road margins. 


364 


JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Lee yA 
\ 


Fics. 74-78. Larval stages of Josia aurifusa. 74, Head, thorax, and segments Al-A3 
of fourth instar, lateral view; 75, Segments A6-A10 of fourth instar, lateral view (scale 
line = 2 mm); 76, Head of fourth instar, frontal view (scale line = 1 mm); 77, Prothoracic 


shield of fourth instar, dorsal view (scale line = 0.5 mm); 78, First instar, lateral view 
(scale line = 1 mm). 


Egg. Diameter = 0.7 to 0.8 mm; found in small clusters of two to nine eggs. Duration: 
4 days. Development time from egg to adult = 42 days. 


First Instar. Similar to other Josiini; head dark brown, slightly lighter on front. Body 
length = 4.05 mm; head width = 0.48 mm (n = 1). 


VOLUME 48, NUMBER 4 365 


Fics. 79-81. Scanning electron micrographs of Josia aurifusa larvae. 79, Ventral 
portion of first instar head, lateral view ( x 350); 80, Right antenna of fourth instar, frontal 
view (xX 266); 81, Distal portion of fourth instar antenna showing segments 2 and 3 ( x 734). 
[An = antennal segment. ] 

Fic. 82. Right tarsus on T2 of fourth instar Josia turgida larva, mesal view (x 292). 


Second and Third Instars. Similar in coloring and setal pattern to final instar. Head 
widths: 2nd instar = 0.81-0.83 mm; 8rd instar = 1.22-1.26 mm (n = 5). 

Final (Fourth) Instar. Head (Fig. 85) white with wide, brown, irregularly patterned 
frontolateral stripes extending from stemmata up to epicranial suture, stripes widening 
near suture; labrum black with a white central region; antenna with segment 2 relatively 
short, less than twice the length of segment 1. Prothoracic shield with an unpigmented 
medial seam, anterolateral angles produced, curled inward (Fig. 86). Ground color of 
thorax and abdomen reddish maroon (Fig. 19); red colored regions lightly spotted with 
white (Figs. 83, 84); white spots present at bases of primary setae; yellow-white dorsal 
and lateral patches larger than in J. aurifusa, margins smoother; venter white, with red 
transverse connecting lines on Al and A2; L3 on A3—A6 surrounded by a large white 
patch; segment AQ conspicuous, entirely white above (Fig. 19). Tarsal setae (Fig. 82) 
similar to Josia gopala. Primary setae (Figs. 83-85) as in J. radians and J. aurifusa; 
lateral plate of Al0 proleg base with approximately 9 setae. Body length = 24.0-25.0 
mm; head width = 1.88-1.94 mm (n = 3). 


366 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fics. 83-86. Larval stages of Josia turgida. 83, Head, thorax, and segments Al—A3 
of fourth instar, lateral view; 84, Segments A6—A10 of fourth instar, lateral view (scale 
line = 2 mm); 85, Head of fourth instar, frontal view (scale line = 1 mm); 86, Prothoracic 
shield of fourth instar, dorsal view (scale line = 0.5 mm). 


Pupa. Cremaster similar to J. aurifusa, slightly less concave ventrally; eight hook- 
shaped setae present. Duration: approximately 17 days. 

Adult. FW length = 14.0-17.0 mm. Head and antenna black; face white with front 
blackish gray; scales at antennal base white; pectinations of male antenna moderately 
long; labial palpus porrect, relatively long and thin, extending well beyond front; basal 
three fourths of segment 1 white, rest of palpus black. Dorsum of thorax black, an orange 
stripe on each side running from patagium to base of abdomen; tegula mostly orange, 
black at base; lateral portion of thorax orange; coxa black on lateral surface, white on 
anterior surface; legs dark gray, tibiae whitish-buff below. FW black with a wide, orange 
longitudinal stripe from base to within 2 mm of outer margin (Figs. 32, 33); stripe tapered, 
slightly irregular along anterior margin near apex; Rs black at base, orange beyond; HW 
(Figs. 32, 33) orange with upper angle black from a point near upper corner of DC to 
outer margin; a diffuse black patch along 2A near outer margin. Abdomen black with 
wide, orange lateral stripes, stripes becoming buff at caudal margin of A8; venter white. 


VOLUME 48, NUMBER 4 367 


Discussion. Josia turgida, described by Warren (1905b:314) in the 
subfamily Cyllopodinae (Geometridae), was considered a form of J. 
aurifusa by Hering (1925) and subsequent authors, but we here elevate 
it to species status. 

Although J. turgida and J. aurifusa are extremely close, we found 
what seem to be reliable differences between these two sympatric spe- 
cies. Adults of J. turgida generally are more orange than those of J. 
aurifusa (compare Figs. 32-86). The FW stripes are wider and the 
black portion of the HW does not reach the wing base. In J. turgida 
there is usually a prominent orange macula on the dorsum of the 
mesothorax, while in J. aurifusa this area is either entirely black, or 
has a very small, faint orange spot. There are also subtle differences 
between the larvae. The yellow-white lateral patches on the thorax and 
abdomen are larger in J. turgida than in J. aurifusa (compare Figs. 
74, 75 with 88, 84), especially those surrounding seta L8 on the proleg- 
bearing segments. The head stripes of J. turgida are lighter in color, 
and the pigmentation is more irregular (Fig. 85). The larvae of J. turgida 
are thus generally lighter in color than those of J. aurifusa. 

Josia aurifusa and J. turgida, together with their various color forms, 
belong in a species complex with Josia auriflua Walker, known from 
the western Andes of Colombia south to Bolivia. To further complicate 
matters, three Dognin names—inaequiflexa, scalata, and flavipars— 
are listed as subspecies of J. auriflua in Bryk (1980). The entire aurifusa 
complex poses a serious taxonomic challenge; wing pattern variation 
seems to merge, and genitalic differences are not always clear-cut (JSM, 
unpubl.). 


CONCLUSIONS 


Based on our findings for these five Venezuelan Josiini and on pub- 
lished reports for other taxa, we now can offer a general summary of 
biological characteristics for the tribe. 

The species occur over a wide range of elevations, from lowland 
jungle to shrubby, high altitude sites in the Andes of South America 
and the Central Cordillera of Central America. Cyanotricha bellona 
has been recorded at 4000 m in Peru (BMNH), and Josia gephyra was 
collected in Colombia at 4700 m (see discussion of J. gopala above). 
Josiini are most common in secondary forests and disturbed habitats, 
but they also occur in pristine environments. Even in primary forests, 
however, they seem to favor light gaps, forest edges, and stream mar- 
gins, perhaps due to the higher frequency of their passifloraceous hosts 
in such habitats. In this study, immatures of Josia radians were found 
on Passiflora growing in a coffee plantation, and at another site close 
to the city of Mérida within the confines of a housing development. 


368 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


At least one species is of economic importance. Cyanotricha necyria 
is an occasional pest in commercial fields of Passiflora mollissima in 
the Andes of Colombia, Ecuador, and Peru (Castaneda 1956, Martin 
& Nakasone 1970). Markin et al. (1989) have studied C. necyria and 
other Josiini as potential biological control agents of P. mollissima, a 
damaging weed in Volcano National Park, Hawaii. 

Members of the Josiini share many life history features, some of 
which apparently are unique to the tribe. For example, unlike most 
Lepidoptera, which have either five or six larval instars, all Josiini so 
far studied have only four (Spitz 1931, d’Almeida 1932, Markin et al. 
1989). The number of larval instars in other subfamilies of the Noto- 
dontidae is five (Packard 1895), and published reports for Dioptinae 
outside the Josiini indicate that five is typical there as well (Herbert 
1920, Wolda & Foster 1978). 

In the species we observed, and in Cyanotricha necyria (Markin et 
al. 1989), first instars often feed together on the same leaf, while sub- 
sequent instars feed individually. These observations agree with pre- 
vious work showing this to be a general trend for the Notodontidae 
(Godfrey et al. 1989). Associated with this trend is a developmental 
change whereby the first instar mandibular margin is serrate, while 
that of later instars is smooth (Godfrey et al. 1989, Dockter 1993). 

Josiini exhibit a characteristic pupation behavior. The pre-pupal cat- 
erpillar makes a shelter, using strong silk threads to weave pieces of 
leaves or debris together. The larva then pupates in this enclosure, 
normally within a few days after its construction. In the absence of 
such materials, the larva builds a flimsy silk net. Pupation usually takes 
place off the hostplant, but in one instance we found a pupa on its 
Passiflora host. 

Several characteristics distinguish the immatures of Josiini from those 
of heliconiines, the only other Lepidoptera common on Passiflora. Hel- 
iconius eggs, deposited singly or occasionally in large clusters, are some- 
what cylindrical with prominent surface sculpturing, and are usually 
yellow, orange, or red (Beebe et al. 1960, Benson et al. 1976, DeVries 
1987). They are generally larger than the eggs of Josiini. Female Hel- 
iconiinae frequently oviposit on new leaves or on shoots near the mer- 
istem, whereas josiines tend to lay on older leaves closer to the ground, 
always on the leaf undersurface. The eggs of Eueides (Heliconiinae) 
are similar to those of Josiini in that they are frequently laid on the 
undersurface of mature leaves, and they are small and green. However, 
Eueides eggs differ in having prominent surface sculpturing (JSM pers. 
obs.). 

Josiine larvae lack projections on the body. They exhibit the sha- 
greened cuticle characteristic of other Dioptinae (Fracker 1915, Forbes 


VOLUME 48, NUMBER 4 369 


1939, Miller 1991), a trait that can be seen with the aid of a hand-lens. 
The caterpillars thus are impossible to confuse with heliconiine larvae, 
which are usually greenish white overlaid with various dark patterns, 
and which have long, spiny projections on the head, thorax, and ab- 
domen (Beebe et al. 1960, Brown 1981, DeVries 1987). 

Adult josiines can be recognized in the field by their erratic, fluttering 
flight and bright coloration. After landing first on the upper surface of 
a leaf, they often move quickly out of sight to the underside. Upon 
close examination, the unusual metathoracic tympanum and wing ve- 
nation can be used as diagnostic features to distinguish them from all 
other Lepidoptera. 

Having here described the immature stages of four Josia species and 
Thirmida discinota, we hope to discover life histories for still other 
taxa. We urge field workers collecting on or around Passiflora to learn 
to recognize and collect Josiini. A long-term goal is to accumulate 
enough hostplant data to compare patterns in the Josiini with those that 
have been described in the literature for the Heliconiinae (Benson et 
al. 1976). Because the immature stages appear to be a rich source of 
characters, their discovery also will benefit greatly systematic studies 
on the Dioptinae. 


ACKNOWLEDGMENTS 


We are grateful to INPARQUES (Instituto Nacional de Parques) for permission to 
collect in Venezuelan national parks. For permission to study at Rancho Grande, we 
thank Alberto Fernandez Badillo, Director of the Estacion Biologica Dr. Alberto Fer- 
nandez Yépez. For help with defining life zones for our study sites, we thank José Remigio 
Guevara. Stephen Tillet provided identifications of Passiflora species. We also thank John 
MacDougal (Missouri Botanical Garden) and Christian Feuillet (Smithsonian Institution) 
for discussions concerning Passiflora identification and classification. Miguel Molinari 
helped with field work and additional Passiflora identifications, and Jorge Avila cared 
for moth cultures. For logistical help we would like to thank Alejandro Grajal and Stuart 
Strahl. David Wagner and Tim McCabe provided helpful comments on the manuscript. 
Cal Snyder kindly took the color photographs of adults. 

We also would like to thank the following people and their institutions for the loan of 
adult material: Martin Honey and David Carter (BMNH); John Rawlins and Chen Young 
(CMNH); Jim Liebherr and Rick Hoebeke (CUIC); Tom Emmel and Andre Surakov 
(Florida State Collection of Arthropods) Julian Donahue (LACM); José Clavijo, Vilma 
Savini, and Luis Joly (MIZA); Allen Young (Milwaukee Public Museum); Robert Poole 
(NMNH); Wolfram Mey (ZMH). This research was supported by National Science Foun- 
dation grant BSR-9106517 to JSM. 


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Received for publication 20 October 1998; revised and accepted 18 June 1994. 


Journal of the Lepidopterists’ Society 
48(4), 1994, 373-380 


A SIMPLE PORTABLE TRAP FOR 
MIGRATING BUTTERFLIES 


THOMAS J. WALKER 


Department of Entomology and Nematology, University of Florida, 
Gainesville, Florida 32611, USA 


AND 


JAMES J. WHITESELL 


Department of Secondary Education, Valdosta State University, 
Valdosta, Georgia 31698, USA 


ABSTRACT. An economical, easily built, 4-m wide trap for migrating butterflies was 
developed and tested. Made of thin-walled electrical conduit, nylon rope, and polypro- 
pylene netting, a “simplex” trap can be erected or taken down quickly, rolled into a 
compact bundle, and carried by one person. 


Additional key words: migration, Phoebis sennae, Agraulis vanillae, Florida. 


Migrating butterflies are peculiarly subject to trapping because they 
generally fly in a straight line near the ground, and upon encountering 
an obstacle, they attempt to fly over it rather than change directions. 
Starting in 1975, a series of traps was developed that took advantage 
of this behavior (Walker 1978, 1985b, Walker & Lenczewski 1989). 
These were used to monitor butterfly migrations in the southeastern 
United States (Walker 1985a, 1991, Walker & Riordan 1981, Lenczews- 
ki 1992). 

Since 1986, one of us (JJ)W) has headed a project to promote the 
study of butterfly migration by high school science students throughout 
Georgia. A major goal of these studies has been to mark and release 
migrating Phoebis sennae (L.) (Pieridae) and Agraulis vanillae (L.) 
(Nymphalidae) in hopes of documenting their migration routes and 
distances. As an adjunct to this project, we worked together to develop 
a trap that was economical, easy-to-build, and effective. We succeeded 
in satisfying these criteria with a “semi-portable” trap that had a rigid 
3.4 x 3 xX 1.8 m frame of lumber and metal tubing (Walker & Whitesell 
1998) but soon discovered that such traps, when used on high school 
grounds, were often vandalized or totally destroyed at night or over 
weekends. We therefore added the criteria that the traps had to be 
easily erected, taken down, and moved in and out of school buildings. 


METHODS 


An earlier trap was portable (Walker & Lenczewski 1989) but erect- 
ing it involved setting and adjusting eight guy ropes and took up to an 
hour. Furthermore, building it required difficult sewing, sheet metal 


374 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


work, and cutting and riveting hardware-cloth cages. We conceived, 
built, and tested a series of much simpler portable traps that shared 
these features: (1) end supports of just two lengths of thin-walled elec- 
trical conduit (EMT), held together by a single eyebolt near the top 
(permitting the ends to scissor shut when the trap was taken down), (2) 
the EMT supports of the trap were connected by 4-m lengths of 3.2 
mm braided nylon rope, including one 2 m above ground level and 
two ~5 cm apart at the eyebolt, (3) the trap was covered with poly- 
propylene netting glued to the EMT frame and around lengths of rope 
with silicon caulk, (4) erecting the trap involved opening and positioning 
the end frames and putting the connecting ropes under tension with a 
guy attached at each end to the eyebolt (Fig. 1A & B), (5) trapped 
butterflies were retained in the triangular duct at the top of the trap 
rather than passing into hardware-cloth holding cages. We called these 
traps “‘simplex”’ traps and built and tested six types. 

Five designs of end frames were tested—three sizes of ““A-frames”’ 
and two sizes of “oblique frames’ (Fig. 1C). For convenience, the 
designs were named by the lengths of the end-frame members expressed 
to the nearest meter. In pilot tests in fall of 1991, the 3x3 and the 3x5 
designs proved inefficient. 

The remaining three designs were compared 17 Sep to 21 Oct 1992 
with each other and with two permanent traps that had been in service 
since 1984 (Walker 1985b, 1991). The 5x5 and 4 x6 traps were covered 
with black, nearly invisible, polypropylene netting (~8 x 10 mm mesh, 
Tenax Ornex SM®). Two 4x4 traps were tested, one with the black 
fabric and one with orange, daylight-fluorescent, polypropylene netting 
(~6 X 6mm mesh, Tenax Plurima®). The four traps were set in random 
order facing NNW ina line beginning at the west end of the permanent 
traps. The order was rerandomized after each quarter of the migratory 
season, with the restriction that each trap would occupy each position 
only once. Traps were serviced every one or two days. Catches by the 
simplex traps were compared by ANOVA. The permanent traps were 
excluded from the ANOVA because there was no control for position 
effects. 


RESULTS 


For neither P. sennae or A. vanillae were the numbers caught in the 
four simplex traps significantly different (P=0.08 and 0.85). For both 
species the greatest total numbers were caught in the 4x4 black trap 
(80 and 32) and the least in the 5x5 trap (83 and 25). The total catch 
per meter of simplex trap was 14.2 and 7.7 (P. sennae and A. vanillae) 
and, of permanent trap, 17.0 and 31.3. 


VOLUME 48, NUMBER 4 3795 


3X3 4X4 5X5 3X5 4X6 
(3.0 X 3.0) (3.8 X 3.8) (4.6 X 4.6) (3.0 X 4.6) (3.8 X 6.1) 


Fic. 1. Portable butterfly trap. A. Frame of simplex trap constructed of EMT (solid 
lines) and braided nylon rope (dashed lines). B. Same trap covered with polypropylene 
netting. C. End frame configurations that were tested, with EMT lengths, in m, below. 
D. Technique for splicing two lengths of EMT. Splint is 40” piece of 4%” EMT with 
wrappings of tape in four places. Splice is made by sliding pieces of %” EMT over splint. 


DISCUSSION 


Of the simplex designs tested, we recommend the 4 x 4 traps because 
they caught as many or more P. sennae and A. vanillae as larger, 
harder-to-build-and-service designs. Using the directions included in 
the appendix of this paper, Georgia high school students and their 
teachers have built and put in service at least 15 4x4 simplex traps. 

The orange polypropylene netting was tested in hopes that it would 
be a magnet to P. sennae, which stops at red flowers (and red taillights 
in parking lots). It actually caught fewer P. sennae and A. vanillae, 
but not significantly fewer. A comparison of the two fabrics on a pair 
of semi-permanent traps likewise failed to reveal a significant difference 


376 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


in the catches of P. sennae, but the orange caught significantly more 
A. vanillae (Walker, unpublished). A disadvantage of the daylight flu- 
orescent orange fabric is that it makes a trap ultra conspicuous to 
passersby. 

The permanent traps are approximately 60 and 35% efficient at 
catching migrating P. sennae and A. vanillae respectively (Walker 
1985b). Assuming that their position relative to the permanent traps 
did not influence the numbers of potential captives or trapping effi- 
ciency, the simplex traps caught an estimated 50% of the P. sennae 
they should have intercepted and 9% of the A. vanillae. These estimates 
are likely low because the simplex traps caught significantly fewer 
migrants when in the two west-most positions than when in the two 
positions closer to the permanent traps. This may have resulted from 
trees to the NNW of the west-most positions causing migrants to fly 
higher as they approached. 

Compared to previous traps for migrating butterflies, simplex traps 
are the least expensive and the easiest to build. They are easy to erect, 
strike, carry, and store. For the first time, a researcher could, in a single 
day, set out an extensive line of traps to quantify a migration, and move 
it to a different transect the next day. Unlike netting used previously, 
the two types of polypropylene that were tested did not weaken after 
more than a year in the Florida sun; and because the traps are easy to 
take down and store, there is no need to leave them outside when not 
in use. Because the width of the trap is determined by ropes under 
tension rather than the standard 3m length of EMT crosspieces, simplex 
traps can take advantage of the full 4 m width of the fabric, increasing 
the reach of the trap by 33%. 

The most serious disadvantage of simplex traps is that they are sub- 
stantially less efficient in catching A. vanillae than the semi-portable 
traps. Lacking a valved catching cage, simplex traps must be serviced 
at 1 or 2-day intervals or captured butterflies will escape. Finally, the 
ease with which they can be taken down means that simplex traps are 
easily stolen (if left up). 


ACKNOWLEDGMENTS 


We are grateful to Evandro de Oliveira, Barbara Lenczewski, and James Lloyd for 
constructively criticizing the manuscript. This paper is Florida Agricultural Experiment 
Station Journal Series No. R-03545. 


LITERATURE CITED 


LENCZEWSKI, B. 1992. Butterfly migration through peninsular Florida. Ph.D. disser- 
tation, University of Florida, Gainesville. 132 pp. 

WALKER, T. J. 1978. Migration and re-migration of butterflies through north peninsular 
Florida: Quantification with Malaise traps. J. Lepid. Soc. 32:178-190. 


VOLUME 48, NUMBER 4 OUN 


. 1985a. Butterfly migration in the boundary layer, pp. 704-728. In M. A. Rankin 

(ed.), Migration: Mechanisms and adaptive significance. Contrib. Marine Sci., suppl. 

vol. 27. 

1985b. Permanent traps for monitoring butterfly migration: tests in Florida, 

1979-84. J. Lepid. Soc. 39:313-320. 

1991. Butterfly migration from and to peninsular Florida. Ecol. Entomol. 16: 
241-252. 

WALKER, T. J. & B. LENCZEWSKI. 1989. An inexpensive portable trap for monitoring 
butterfly migration. J. Lepid. Soc. 43:289-298. 

WALKER, T. J. & A. J. RIORDAN. 1981. Butterfly migration: are synoptic-scale wind 
systems important? Ecol. Entomol. 6:433-440. 

WALKER, T. J. & J. J. WHITESELL. 1993. A superior trap for migrating butterflies. J. 
Lepid. Soc. 47:140-149. 


Received for publication 8 December 1998; revised and accepted 8 February 1994. 


APPENDIX: BUILDING AND USING A SIMPLEX TRAP 


Materials 


5 — 10’ pcs, ¥” EMT (electrical conduit) 

2 — 10’ pc, 4%” EMT [will have 40” pc left over] 

8 — 1 ¥,” x '°/,, machine screws with nuts 

2 — ¥,” eyebolts 3” long with nuts 

6 — Y¥,” machine washers 

2— Y¥,” x 1 Y,” machine bolts with nuts 

2 tubes transparent silicon caulk 

492' of 13’-wide, %%”-mesh, [filament dia = 0.15 mm], black polypropylene 
netting. (“Tenax Ornex SM”: Geotenax Corp., Jessup MD) 

118%’ of 4%” braided nylon rope 

AO’ of *%,” braided nylon rope 

1 roll electrical or other tape 

1 roll flag tape 


Tools and Reusable Supplies 


1 Ib. 8d box nails 

1 8’ step ladder 

1 caulk gun 

1 heavy hammer 

1 tape measure 

1 nail apron 

1 pr. scissors 

1 drill with 4,” and '%,” bits 


Procedures 


Prepare EMT, ropes, and stakes. Cut one of the lengths of 4” EMT 
into four 2.5’ pieces. Cut one of the lengths of 4” EMT into three 40” 


378 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


pieces; from the other, cut one 40” piece and two 20” pieces. Cut the 
’z’ rope into seven 13.5’ pieces and two 12’ pieces. Cut the ¥%,” rope 
into two 20’ pieces. Use a lighter or blow torch to melt the ends of the 
'*" ropes. Pull each melted end to a point to aid in threading the rope 
through snug holes. (Warning! Melted nylon is hot and sticky. Protect 
your fingers by using leather gloves or thick layers of cloth.) Melt the 
ends of the *%,” ropes to prevent unraveling. To finish the stakes, drill 
a Y,”-dia hole about 1” from one end of each 20” piece of 4” EMT. 
Insert and fasten a 1 ¥,” machine bolt in each hole—to prevent the guy 
ropes from slipping off the tops of the stakes. 

Make support poles. Make 12.5’ support poles by joining 2.5’ pieces 
of ¥%,” EMT with 10’ pieces. The segments are joined by an internal 
splint made of a 40” section of 4” EMT. To prepare the splints, wrap 
the four 40” pieces of 4” EMT with tape at each end and at 1” on 
either side of the midpoint (Fig. 1D, splint). At each of the four wrap- 
ping places on each piece the tape should increase the diameter of the 
EMT so that it will slide snugly into both to-be-joined pieces of %/” 
EMT. Join each 2.5’ piece of *,” EMT to one of the 10’ pieces by slipping 
a wrapped 40” piece of 4” EMT 20” into the two pieces to be joined 
(Fig. 1D, splice). Drill a 1%,” dia hole through the assembly at 1” on 
each side of the junction of the pieces of °/,” EMT. Secure with two 14” 
x 10-32 machine screws. Designate two of the support poles as eves 
poles and two as rear poles. Make the 2.5’ segment the upper end of 
each pole. 

Measuring from the top of each eves pole, drill a ¥,”-dia hole at 12”. 
In the same plane, drill !°4,”-dia holes at 0.5, 18, 57, and 149.5” (=0.5” 
from bottom) (these are for the top, slot, eves, and bottom ropes). Drill, 
at 90° to the other holes, a '%,”’-dia hole Y,” from the bottom (for the 
end rope). Drill the rear support poles like the eves poles, but omit the 
hole for the eves rope (i.e., the hole 57” from the top). 

Assemble and erect the frame. Insert a 3” eye bolt through the Y,” 
holes to join the eves and rear poles of each end of the frame. Place a 
washer at the eye, between the poles, and next to the nut. The eyes 
establish the outside of each end, and one end must have the eves pole 
on the outside and the other must have it on the inside. Lay the ends 
on the ground several feet apart, one with outside (=eye-side) up and 
the other with outside down. Slightly spread the poles of each end 
making the front and rear poles of each end parallel to the corresponding 
pole of the other. Now thread the seven 18.5’ pieces of rope through 
corresponding holes of the two ends being careful not to cross the ropes 
and to thread the ropes from the inside of the ends. To secure the ropes 
to the support poles, tie an overhand knot in each end, leaving a tail 
of about 1”. Thread the two end ropes into the bottommost holes and 


VOLUME 48, NUMBER 4 379 


secure them in the same way as the cross ropes. Attach a %,” guy rope 
to each eye. 

Drive the two stakes into the ground ~40’ apart. With one person 
on each end, erect the support poles with eyebolt eyes outward. Open 
the end poles into A’s, being careful to face the two eves poles in the 
same direction. Tie the guy ropes to the stakes. Check the cross ropes 
and end ropes for problems—viz., wrong insertions and crossed ropes. 
Undo, rethread, and retie as required. Use guy ropes to pull cross ropes 
taunt and move poles to tighten end ropes and to square the frame in 
all its planes. 

Apply the netting. Cut the netting into one 19’ and two 11.5’ lengths. 
Starting at the bottom rear of the trap, thread the 19’ piece up between 
the slot ropes, outside and over the top ropes, and back down between 
the slot ropes. Once threaded, spread the netting laterally, and starting 
at the lower rear corners temporarily attach it around the frame, to 
itself, using 8d box nails as you would straight pins in cloth. Pull the 
netting around the bottom cross rope making a 1” hem. At intervals of 
ca. 6” insert two nails parallel to the rope and %” and ¥,” from it. Pull 
the netting around each rear pole and insert nails at ca. 6” intervals. 
Continue to the top of the trap and then down the eves pole. Upon 
reaching the eves, hem the netting around the eves rope in the manner 
described for the bottom rope. Now put a bead of silicon caulk between 
each pair of nails in the two hems, being careful to ensnare both layers 
of netting. 

Wrap the selvage of an 11.5’ piece of netting around each end rope 
and use nails to hem as before. Temporarily attach the middle of the 
opposite selvage to the eye near the top of the trap. Now work from 
the bottom and stretch the netting between eves pole and rear pole. 
Use nails to tack it to the main net and around the lower portion of 
the eves pole. Make your work easier by cutting off surplus netting (as 
it becomes apparent what is surplus). Once you have the netting on 
each end loosely fit and trimmed, remove and reinsert the nails as you 
pull it tight for the final fit. When you have all the netting in place 
and stretched except for two small triangles that close the ends of the 
duct, glue the netting to the support poles with a bead of silicon running 
the full length of each pole. When the silicon has congealed, remove 
all nails. 

Take two scraps of netting and use nails to sew them to each end of 
the duct. Loosely trim the netting and stretch the triangles so that they 
can be glued with silicon to the poles on either side and to the selvage 
of the main net above. When you apply the silicon be sure to close any 
holes (=potential butterfly exits) that remain where the main net changes 
from one support pole to another. When the silicon has congealed, 


380 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


remove the nails. Then check again for holes larger than the 4” mesh 
of the netting and close them with silicon. 

To make the netting visible to persons working around the trap, tie 
1’ lengths of flag tape to the netting at 2’ intervals at 4’ above the 
ground. 

Striking, transporting, erecting, and servicing the trap. To strike a 
trap, a person at each stake unties the guy rope and keeps it taunt as 
he/she walks toward the trap. Upon reaching the trap, each grabs a 
rear pole and tips the top forward causing the rear and eves poles to 
scissor shut while making sure the end netting tucks inside the closing 
poles. The support poles are then held horizontally with the main net 
stretched between and the guy ropes thrown toward the center. The 
two strikers walk toward each other as each rolls the netting around 
two folded poles. When the rolls meet near the center, one takes both 
rolls. The struck trap can now be transported on one person’s shoulder. 
The rolls can be secured with a bungie cord or rope. 

To erect a trap, two persons reverse the striking process, being careful 
not to tear the netting as it unrolls. Unless the end of the eye bolts and 
the splint-securing machine screws have been cut off and filed, the 
netting may have to be extricated from them. Drive stakes 45’ apart 
along a line that is perpendicular to the mean direction of migration. 
If there are not two bowlines in each guy rope, tie them now—first 
one to make the eye-to-stake length 18.5’ and then one to make that 
length 24’. Now erect the trap making sure that it is square, centered 
between the stakes, and facing into the migratory stream. With the 
inner bowline of each guy rope around a stake, the trap’s cross ropes 
should be properly taunt. If they are not, adjust one of the bowlines. 

To remove butterflies from the trap, partially collapse it by releasing 
the guy on one end until its outer bowline is holding the trap. Butterflies 
in the two thirds of the duct closest to that guy should now be easily 
reached. Restore the trap and collapse it in the other direction to reach 
butterflies in the rest of the duct. 


GENERAL NOTES 


Journal of the Lepidopterists’ Society 
48(4), 1994, 381-386 


STICKY INTEGUMENTAL COATING OF A DALCERID CATERPILLAR: 
A DETERRENT TO ANTS 


Additional key words: Chemical defense, Dalcerides ingenita, Lepidoptera, larva, 
Camponotus floridanus. 


Dalceridae are a small group (85 species) of mostly neotropical moths with sluglike 
larvae covered with a gelatinous coating (Dyar 1925, Hopp 1928, Stehr & McFarland 
1987, Miller 1994). Nothing was known about the chemistry or function of this coating, 
although it had been proposed to serve for defense (Stehr & McFarland 1987). We here 
present evidence, based on staged encounters between a dalcerid caterpillar and ants, 
that the larval coating can indeed serve as an effective deterrent to attack. 

The larvae were offspring of gravid females of Dalcerides ingenita (Hy. Edwards), 
the only species of Dalceridae native to the United States, collected at Ash Canyon in the 
Huachuca Mountains, Cochise County, Arizona. They were raised on mature foliage of 
one of their host plants, Arctostaphylos pungens (H.B.K.) (Ericaceae), as well as on leaves 
of various oaks [Quercus emoryi Torrey (Fagaceae) figures in the moth’s natural diet]. 
Voucher specimens of adults and larvae are deposited in the entomological collection of 
the National Museum of Natural History, Washington, D.C. 

The dorsal investiture of D. ingenita has the warty appearance (Fig. 1) characteristic 
of dalcerid larvae (Dyar 1925, Stehr & McFarland 1987). The investiture is moderately 
sticky and can be readily pulled from a larva by rolling it onto a glass rod (Fig. 2). Pulling 
on a single wart with forceps typically results in detachment of a string or cluster of warts 
(Fig. 2). If a larva is artificially denuded by removal of the entire complement of warts, 
a coating of semi-liquid material is exposed (Fig. 1). 

The ants used in the encounters were from a laboratory colony of Camponotus flori- 
danus (Buckley) (Formicidae), originally taken near Lake Placid, Florida. We previously 
had used this species in predation experiments with other insects (Eisner 1972). 

The tests were carried out in 15 x 50 mm glass petri dishes. For each test, a number 
of ants (5-10) were first added to a dish, following which a piece of Quercus leaf was 
introduced bearing a single D. ingenita larva. Events were monitored with a Wild M400 
photomicroscope and were consistent for each of the 5 larvae tested. Ants repeatedly 
came in contact with the larvae and inspected these with antennae and palps, but in most 
cases they withdrew without attempting to bite. There was no evidence of repellency on 
near contact. Ants seemed to back away from larvae only after directly palpating them. 
Actual bites or attempted bites occurred only in a few cases, but the results in these 
instances were dramatic. Ants either became temporarily stuck to the larval coating and 
had to struggle briefly to free themselves, or they pulled away quickly, but with their 
mouthparts encumbered by detached dabs of coating (Fig. 3). Such ants never persisted 
in their assault, but engaged instead in protracted, eventually successful, cleansing activ- 
ities. 

Two samples of coating, each representing strippings from several larvae, were extracted 
respectively with methanol and ether, and examined by gas chromatography (using a 
non-polar liquid phase). No evidence was obtained of presence of volatile or volatilizable 
components. Given that the coating appeared to effect its deterrency by mechanical action 
rather than repellency, this finding was perhaps to be expected. 

Caterpillars possess a multiplicity of chemical defenses, including eversible (Brower 
1989) and dischargeable glands (Attygalle et al. 1993), urticating spines (Kawamoto & 
Kumada 1984), and glandular hairs (R. Rutowski & T. Eisner personal observation, 
Eurema and Pieris). In addition, some derive protection from forcible emission of enteric 


382 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fic. 1. Typical larva, showing rows of dorsal gelatinous warts (top). Larva from which 
middorsal warts have been removed, exposing the semi-liquid material beneath (bottom). 
[Scale bar = 2 mm] 


fluid (Smedley et al. 1993) or possession of systemic toxins acquired from the diet (Bowers 
1993). To our knowledge, dalcerid larvae are exceptional among caterpillars in possessing 
a sticky integumental coating, essentially comparable to that of slugs (Mollusca: Gastrop- 
oda). Olona spp. of the family Limacodidae, a group related to the Dalceridae, also possess 
a gelatinous coating (Holloway et al. 1987). The integument of molluscan slugs has itself 
been shown to be deterrent to ants (Eisner 1970). Sticky integumental coatings are 
generally rare among insects. They occur in certain sawfly larvae (Caliroa spp., Ten- 
thredinidae), in which they also may be defensive, although there appears to be no 
evidence to that effect (Nordin et al. 1972). In some cockroaches, the last abdominal 


VOLUME 48, NUMBER 4 383 


Fic. 2. A wart’ has been grasped with forceps and is being pulled away, causing a 
strand of warts to be detached from the investiture (top). A cluster of warts has been 
taken up by a glass rod; to remove the cluster the rod was pushed into the investiture 
and slowly rolled (bottom). [Scale bars: top = 1 mm; bottom = 2 mm] 


tergites are covered with a proteinaceous slime of proven physical deterrency to ants 
(Plattner et al. 1972). A number of arthropods produce sticky secretions as products of 
dischargeable glands. Examples include certain syrphid fly larvae, which protect them- 
selves by use of a viscous glue that they discharge from salivary glands (Eisner 1972), 
and geophilid centipedes, which eject a proteinaceous cyanogenic slime when disturbed 
(Jones et al. 1976). A classic example is that of onychophorans (Peripatus and its relatives), 


384 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Fic. 3. Camponotus floridanus biting a larva; the left mandible has become entangled 
in the sticky slime (top). Camponotus floridanus backing away after having bitten into 
the larva’s investiture; a wad of slime has become stuck to the ant’s mouthparts (bottom). 
[Scale bar = 2 mm] 


which eject aimed jets of a rapidly hardening glue from a pair of cephalic glands (Al- 
exander 1957, Eisner 1970). The fluid literally cements attacking ants and spiders to the 
substrate when these are hit by the spray (Eisner, unpublished laboratory observations 
with unidentified onychophorans from Panama). 

Insufficient D. ingenita were available to determine precisely how the integumental 
coating is engendered. Evidence presented so far about the presumed underlying secretory 


VOLUME 48, NUMBER 4 385 


mechanism suggests that the coating may consist in part of sloughed integumental lay- 
erings (Hopp 1928). 

This is paper no. 123 of the series Defense Mechanisms of Arthropods; no. 122 is 
McCormick, K. D. et al. (in press), Tetrahedron. This study was supported in part by 
grant AI02908 from NIH. We thank Maria Eisner for preparation of the illustrations, 
Athula Attygalle for the chromatographic data, and Noel McFarland for providing the 
gravid moths. The ant colony was collected at the Archbold Biological Station, Lake 
Placid, Highlands County, Florida. 


LITERATURE CITED 


ALEXANDER, A. J. 1957. Notes on onychophoran behavior. Ann. Natal Mus. 14:35-48. 

ATTYGALLE, A. B., S. R. SMEDLEY, J. MEINWALD & T. EISNER. 1993. Defensive secretion 
of two notodontid caterpillars (Schizura unicornis, S. badia). J. Chem. Ecol. 19:2089- 
2104. 

Bowers, M. D. 1993. Aposematic caterpillars: Life-styles of the warningly colored and 
unpalatable, pp. 331-371. In Stamp, N. E. & T. M. Casey (eds.), Caterpillars: Eco- 
logical and evolutionary constraints on foraging behavior. Chapman & Hall, New 
York. 

BROWER, L. P. 1989. Chemical defense in butterflies, pp. 109-134. In Vane-Wright, 
R. I. & P. R. Ackery (eds.), The biology of butterflies. Princeton Univ. Press, Princeton, 
New Jersey. 

Dyar, H. G. 1925. A note on the larvae of the Dalceridae (Lepidoptera). Insec. Inscit. 
Menst. 13:44—47. 

EISNER, T. 1970. Chemical defense against predation in arthropods, pp. 157-217. In 
Sondheimer, E. & J. B. Simeone (eds.), Chemical ecology. Academic Press, Inc., New 
York. 

1972. Chemical ecology: On arthropods and how they live as chemists. Verh. 
Deutsch. Zool. Gesellsch. 65:123-137. 

HoLLoway, J. D., M. J. W. Cock & R. D. DE CHENON. 1987. Systematic account of 
south-east Asian pest Limacodidae, pp. 15-117. In Cock, M. J. W., H. C. J. Godfray 
& J. D. Holloway (eds.), Slug and nettle caterpillars: The biology, taxonomy, and 
control of the Limacodidae of economic importance on palms in south-east Asia. 
CAB International, Wallingford. 

Hopp, W. 1928. Beitrag zur Kenntnis der Dalceriden. Deutsche Entomol. Zeits. Iris 42: 
283-287. 

JONES, T. H., W. E. CONNER, J. MEINWALD, H. E. EISNER & T. EISNER. 1976. Benzoyl 
cyanide and mandelonitrile in the cyanogenetic secretion of a centipede. J. Chem. 
Ecol. 2:421-429. 

KAWAMOTO, F. & N. KUMADA. 1984. Biology and venoms of Lepidoptera, pp. 291- 
330. In Tu, A. T. (ed.), Handbook of natural toxins, Vol. 2, Insect poisons, allergens, 
and other invertebrate venoms. Dekker, New York. 

MILLER, S. E. 1994. Systematics of the neotropical moth family Dalceridae (Lepidop- 
tera). Bull. Mus. Comp. Zool. 153. 

NORDIN, G. L. & E. L. JOHNSON. 1984. Biology of Caliroa quercuscoccineae (Dyar) 
(Hymenoptera: Tenthredinidae) in central Kentucky. II]. Development and behavior. 
J. Kansas Entomol. Soc. 57:569-579. 

PLATTNER, H., M. SALPETER, J. E. CARREL & T. EISNER. 1972. Struktur und Funktion 
des Driisenepithels der postabdominalen Tergite von Blatta orientalis. Zeitschr. Zell- 
forsch. 125:45-87. 

SMEDLEY, S. R., E. EHRHARDT & T. EISNER. 1993. Defensive regurgitation by a noctuid 
moth larva (Litoprosopus futilis). Psyche 100:209-221. 

STEHR, F. W. & N. MCFARLAND. 1987. Dalceridae (Zygaenoidea), pp. 460-462. In 
Stehr, F. W. (ed.), Immature insects. Kendall/Hunt, Dubuque, Iowa. 


Marc E. EPSTEIN, Department of Entomology, National Museum of Natural History, 
Smithsonian Institution, Washington, District of Columbia 20560, USA; SCOTT R. 


386 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


SMEDLEY AND THOMAS EISNER, Section of Neurobiology and Behavior, Cornell Uni- 
versity, Ithaca, New York 14858, USA. 


Received for publication 28 December 1998; revised and accepted 10 April 1994. 


Journal of the Lepidopterists’ Society 
48(4), 1994, 386-388 


HOST PLANTS OF POANES MELANE (HESPERIIDAE) 


Additional key words: grasses, Poaceae, skipper, biology. 


Poanes melane (Edwards), the umber skipper [formerly Paratrytone melane (Burns 
1992)], is found along the Coastal Range of California and in the foothills of the Sierra 
Nevada. In rural areas adults are found in grassy habitats along streams and in forests 
(Emmel & Emmel 1973). This species is common in urban and suburban areas, most 
likely because its host plants (grasses; Poaceae) are abundant and well-maintained in lawns 
(Heppner 1972, Brown 1984). This source of host plants could become especially important 
during the summer generation, a dry season in California when many grasses are no 
longer green. Poanes melane larvae are known to feed on several C, and C, grasses, 
including Cynodon dactylon (L.) Pers. (C,), Deschampsia caespitosa (L.) Beauv. (Cj), 
Lamarckia aurea L. (Moench) (C,), and Stenotaphrum secundatum Kuntze (C,), and 
one sedge (Cyperaceae), Carex spissa Bailey (Brown 1984, Scott 1986). 

Adult female P. melane lay eggs singly on the undersurfaces of grass blades. The 
mature larva is approximately 30 mm long with a brown head and dusky yellow-green 
or tan body mottled with black punctations of varying sizes (Comstock & Dammers 1931, 
Emmel & Emmel 1978). Larvae have a blackish mid-dorsal line, a cream lateral stripe, 
and three dark lines on each side (Scott 1976). Larvae also construct tubular shelters by 
tying two or more leaves of their host plants together with silk or by rolling single broad 
leaves into tubes secured with silk along their edges. Mature larvae feed nocturnally, 
generally at the distal ends of tied or rolled leaves. This association between shelter 
formation and feeding activity was used in several cases in this study to determine the 
use of host plants by P. melane. I report here several new host records from Berkeley, 
California made during a four year study of the relative performance of P. melane larvae 
on C, and C, grasses. All plants used in feeding experiments were grown from seed 
available commercially or from the U.S. Department of Agriculture Soil Conservation 
Service, unless otherwise noted. 

Poanes melane larvae were observed feeding on a hedge of bamboo, Phyllostachys 
bambusoides Sieb. and Zucc. (Poaceae, C,), for three generations in 1985 and 1986. 
Complete development to adults was inferred from the presence of pupal cases in larval 
shelters. Oviposition by adults on P. bambusoides was observed on several occasions in 
the field. 

Ehrharta erecta Lam. (Poaceae, C,) is commonly used by larvae in the field (observed 
from 1985 to 1989). In laboratory experiments, P. melane grew well on E. erecta grown 
from seed collected in the field (Barbehenn and Bernays 1992). Oviposition was observed 
on several occasions on E. erecta in the field. 

Late-instar larvae were observed feeding on Lolium multiflorum Lam. (Poaceae, C;) 
on two occasions in 1985. Adult P. melane (one male and one female) were reared from 
these larvae. Lolium multiflorum also supported growth to adulthood in laboratory ex- 
periments (Barbehenn and Bernays 1992), and adults in 0.5 m° cages readily oviposited 
on potted plants. However, in two subsequent years larvae fed one-month-old L. mul- 
tiflor'um commonly rejected it or were unable to grow. 


VOLUME 48, NUMBER 4 387 


Poanes melane larvae (n=4) were collected on Paspalum dilatatum Poir. (Poaceae, 
C,) in March 1985 (Elizabeth Bernays pers. comm.). Larvae were successfully reared on 
Paspalum dilatatum in several laboratory experiments (Barbehenn and Bernays 1992). 

Pennisetum clandestinum Hochst. ex Chiov. (Poaceae, C,) was commonly oviposited 
on in the field. In the laboratory this coarse grass supported good larval growth (Barbehenn 
and Bernays 1992). Pennisetum clandestinum was grown from plants collected in the 
field. 

Sorghum sudanense (Piper) Stapf. (Poaceae, C,), planted in a mixed stand with C. 
dactylon, P. clandestinum, and E. erecta, was commonly oviposited on by adults flying 
in a 13X7X3 m outdoor screen cage in 1986 and 1987. Larvae grew rapidly on S. 
sudanense in laboratory experiments (Barbehenn and Bernays 1992). 

One case was observed in February 1987 of a mature larva feeding on Hordeum 
leporinum (Link) (Poaceae, C,). Later efforts to rear P. melane on this species were 
unsuccessful, however. Fourth-instar larvae commonly rejected H. leporinum leaves from 
plants grown from seed collected in the field. 

Digitaria sanguinalis (L.) Scop. (Poaceae, C,) was found to be an acceptable host plant, 
both for oviposition in the laboratory and for larval growth (Herbert Baker pers. comm. ). 
The source of these plants was not determined. 

Scott’s (1986) listing of Bromus carinatus as a host plant of P. melane is based on a 
field observation by John Lane of oviposition on this species (Langston 1980). Two ad- 
ditional records reported here confirm that B. carinatus is used by P. melane adults and 
larvae. Several mature larvae were found on this species in March 1985 (Junji Hamai 
pers. comm.) and a female was observed ovipositing on B. carinatus in the Berkeley Hills 
in May 1985 (John Brown pers. comm. ). 

In addition to the above observations of host use by P. melane, the ability of larvae to 
use several novel grass species was tested. Larvae were reared on C. dactylon through 
three instars and then fed the following grasses during the fourth instar: Dactylis glomerata 
L. (potted plants from the field), Agrostis palustris Huds., Festuca myuros L., Festuca 
rubra L., and Agropyron cristatum (L.) (all C; Poaceae). Larvae were successfully reared 
on each of these species (Barbehenn and Bernays 1992). Finally, Poa pratensis (L.) 
(Poaceae, C;) was readily accepted by larvae, but it was not determined whether larvae 
develop successfully on this species. 

In addition to certain plantings of L. multiflorum and H. leporinum, three grasses 
were found to be unsuitable as hosts for P. melane: Panicum virgatum (L.), Panicum 
amarulum (Hitch. & Chase), and a native coastal grass Muhlenbergia rigens (Benth.) 
Hitch. (all C, Poaceae). Larvae (n = 3 per species) consistently rejected the Panicum 
species. Muhlenbergia rigens was readily accepted by larvae (n = 15) but did not support 
growth. The unusually tough and thick leaves of M. rigens is a possible factor limiting 
its use by P. melane. 

With the exception of D. caespitosa and B. carinatus, all of the grasses previously 
reported as host plants for P. melane are introduced species. In addition, all of the new 
host plants reported here are introduced species. As noted by Brown (1984), few native 
host plants of P. melane are known. Given the acceptability of a wide variety of grasses 
as host plants, it seems likely that P. melane originally fed on a variety of native grasses 
before the spread of introduced grasses in California. The acceptability of a wide variety 
of grasses by P. melane also suggests that the “indiscriminate” oviposition on grasses by 
captive females (Comstock & Dammers 1981) and a free-flying female (Brown 1984) 
may represent a viable egg-laying strategy. 

Support for this work was provided by NSF grant BSR-8701000 to E. A. Bernays and 
R. V. Barbehenn. C. Don MacNeill and John W. Brown provided helpful information 
on P. melane. I thank C. Don MacNeill and James A. Scott for critical comments on the 
manuscript. 


LITERATURE CITED 


BARBEHENN, R. V. & E. A. BERNAYS. 1992. Relative nutritional quality of C, and C, 
grasses for a graminivorous lepidopteran, Paratrytone melane (Hesperiidae). Oec- 
ologia 92:97-103. 


388 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


BROWN, J. W. 1984. Host records for Paratrytone melane (Edwards) (Hesperiidae). J. 
Lepid. Soc. 38:138. 

BURNS, J. M. 1992. Genitalic recasting of Poanes and Paratrytone (Hesperiidae). J. 
Lepid. Soc. 46:1-23. 

COMSTOCK, J. A. & C. M. DAMMERS. 1931. Notes on the life history of Poanes melane 
Edw. (Lepid.). Bull. South. Calif. Acad. Sci. 30:20-22. 

EMMEL, T. C. & J. F. EMMEL. 1973. Butterflies of southern California. Nat. Hist. Mus. 
Los Angeles Co., Sci. Ser. 26:1-148. 

HEPPNER, J. B. 1972. The distribution of Paratrytone melane and its spread into San 
Diego County (Hesperiidae). J. Res. Lepid. 10:287-300. 

LANGSTON, R. 1980. Season Summary. News Lepid. Soc. 1980(2):14. 

ScoTT, J. A. 1986. The butterflies of North America. Stanford Univ. Press, Stanford, 
CA. 583 pp. 


RAYMOND V. BARBEHENN, Department of Entomological Sciences, University of 
California, Berkeley, California 94720, USA; current address: Department of Biology, 
University of Michigan, Ann Arbor, Michigan 48109-1048, USA. 


Received for publication 10 December 1993; revised and accepted 25 June 1994. 


Journal of the Lepidopterists’ Society 
48(4), 1994, 388-393 


COMMENTS ON THE NATURE AND ORIGINS OF MIGRATIONS OF 
LEPIDOPTERA TO BERMUDA 


Additional key words: evolution, migratory cues. 


“The Lepidoptera of Bermuda” (Ferguson, Hilburn & Wright 1991), which I recently 
reviewed for this Journal (Gaskin 1993), includes an interesting essay in the Appendix 
which, for brevity, I refer to below as Ferguson (1991). In this, he discussed a number 
of problems and paradoxes concerning the nature and origins of the long-distance mi- 
grations of Lepidoptera to Bermuda, stimulating the additional thoughts and comments 
presented in this note. 

I will start by summarizing the general findings of Ferguson et al. (1991): Bermuda is 
an oceanic archipelago, dominated by one large island, that caps a seamount with no 
earlier geological connections to the North American mainland. This archipelago supports 
a disparate, super-saturated lepidopterous fauna, assembled by over-water dispersal at 
various times during the last five hundred thousand years, from southeastern North 
America and the northeastern Caribbean. A small number of endemic species are rec- 
ognized, derived from ancestors in these two regions. The remaining members of the 
Bermudian fauna are morphologically and probably genetically, similar to their source 
populations. 

The biomass, diversity, and composition of the Bermudian fauna must have undergone 
dramatic increases and reductions during the Quaternary concomitant with several radical 
fluctuations in the surface area of the archipelago. 

It was not the primary purpose of Ferguson and his co-workers to develop hypotheses 
concerning the role that interactions of environmental stimuli, physiology, and behavior 
might have played in the evolution of long distance migrations of Lepidoptera to Bermuda. 
Certainly, the phenomenon appears to pose problems for the evolutionary geneticist. If 
there is no demonstrable return flight, then there seems to be no way for the trait to be 
fed back into the source population, and no cumulative selection from one generation to 
the next (Ehrlich 1984). Some of the Lepidoptera which migrate frequently to Bermuda 
have near-global ranges, and are found also on other oceanic islands. Others have close 
relatives in widely distant regions with similar migratory behavior. Ferguson estimates 


VOLUME 48, NUMBER 4 389 


that the migrants to Bermuda make nonstop flights of more than 1000 km, often using 
lower altitude elements of the jet stream. A number of species from the southeastern 
United States found on Bermuda also make well-documented annual migrations into the 
northern states and Canada. Overwintering is unlikely in such cases because these species 
do not exhibit diapause. Ferguson speculates that many of these make return southern 
flights within continental North America that have not yet been detected. Presently the 
best known two-way migration within North America of course, is that of the monarch 
butterfly (Danaus plexippus L.; Nymphalidae). The movements have been copiously 
documented and the basic elements of the migration have been known for many years 
(Urquhart and Urquhart 1978). Northward spring flights in North America by species of 
Vanessa are also well known (Scott 1986). In 1992, for example, I counted about 125 V. 
cardui and V. virginiensis moving through one small part of Guelph, Ontario, Canada 
within about 4-5 hours on 26 April, pausing to nectar at clumps of spring dandelions on 
small patches of waste ground. Weaker southward return flights by V. cardui have been 
observed during late summer and fall in various parts of the United States (Scott 1986). 
There are also recent reports of consistent southerly autumn movements of V. atalanta 
in the United Kingdom (Riley & Riley 1992). Naturally, these observations may reflect 
only the prevailing wind direction at the time. Without resorting to mass alar tagging, 
it is exceedingly difficult to prove conclusively that observed movements of butterflies 
are in fact part of a large-scale migration rather than periodic local activity. It is, alas, 
quite unlikely that the kind of observer effort generated for the monarch butterfly pro- 
grams could be duplicated for large-scale tagging of cutworm moths. 

Recent advances in methods to identify and trace specific combinations of isotopes 
through food chains may give us an additional indirect way of detecting long-distance 
migrations. Systematic sampling of mtDNA and nDNA in given species at different 
localities also may provide insight into population structure and movements. Unfortu- 
nately, absolute methods for satisfactorily distinguishing sampled entities at the population 
level have yet to be perfected for animal species, despite great success in individual DNA 
“fingerprinting.” 

There are other ways of viewing the problems posed by Ferguson, however. It is rarely 
clear what proportion of any insect population migrates. We probably have more data 
on the migration of D. plexippus than for any other butterfly or moth, but some aspects 
are still unclear. Although huge numbers of monarch butterflies migrate from the southern 
wintering grounds into the northern United States and Canada in normal years, some 
still can be found even in southwestern Texas in the summer months. It seems unlikely 
that the migratory instinct is based on an “all or nothing” response in an entire population 
(Ehrlich 1984). Obviously long-distance return migrations, especially when two genera- 
tions are involved as in the case of the monarch, demand the transfer of quite sophisticated 
cue recognition and navigational programming (Baker 1984, Douglas 1986). We have no 
clear idea of the time scale necessary for such developments. 

Ferguson's second puzzle is that the monarch butterflies of Bermuda are now residential, 
and no longer migrate. The migratory impulse presumably can be over-ridden in some 
way by internal systems that can recognize specific environmental cues or stimuli of 
immediate importance to the organism, attention to which will benefit the individual in 
the short-term and increase its chances of passing on its genetic heritage to offspring. 
Ignoring for a moment the fact that Bermudian monarchs are on an oceanic island, the 
question can be seen as part of the more general one “why don't colonial butterflies 
migrate?’ Presumably, the majority tends to remain in the area through repeated exposure 
to local cues, such as the presence of specific microclimatic conditions or food plants that 
are absent for some distance around the colony. Statistically, relatively weak fliers carrying 
out short-distance search movements (random walks) will usually end up back in the 
locality where the cues are strongest or most repetitive. It is quite normal, however, for 
some individuals to disperse up to tens of kilometers from a colony. This happens in the 
case of the European large copper, Lycaena dispar (f. batavus), for example (Pullin et 
al. 1993). If such wanderers fail to find their way back and the colony is effectively 
isolated by unfavorable habitat, then this behavior could potentially threaten the viability 
of the colony. In other seasons the same behavior might enhance its survival by reducing 


390 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


pressure on limited food resources. Under normal circumstances, however, in the absence 
of significant vagility (capacity for dispersal at least to the nearest suitable habitat site), 
local colonies of a few hundred individuals are certain to be extirpated in the long run 
by environmental events, often simple density-independent occurrences such as flood, 
drought, or unseasonable hard frost at some critical point in the life cycle. Stochastic 
events probably play a greater role in colony extinctions than inbreeding. Dempster (1989) 
argued that vagility, as a basic attribute, is vital for survival even for butterflies well- 
adapted to highly specific habitats, because unless isolated colonies are “topped up” at 
intervals by surplus production from adjacent neighboring units experiencing “good 
years, they ultimately will disappear. Colonial butterflies in many parts of North America 
and Western and Eastern Europe have undergone dramatic declines in recent decades 
(Kudrna 1986). The endangered or threatened species are usually those with rather low 
vagility, often staying close to localized food plants. This was not such a problem before 
the advent of drastic modifications to northern temperate ecosystems by the activities of 
humanity, which have greatly reduced and fragmented previously widespread habitats. 
Of course, there always were extirpations of colonies by natural processes, with eradications 
of whole systems during the cold Pleistocene stadials, and some recolonizations in the 
mild interstadials (Dennis 1976). In the case of species with low vagility, recolonizations 
of areas were simply a function of expansion of a particular habitat or biotope. Never- 
theless, the greater the innate vagility of a butterfly species, the greater the chances of 
recolonizations, recovery in numbers, and increased opportunities for individuals to per- 
petuate their genes. Dempster (1989) argued that it was no accident that the butterflies 
which did not seem to be on the decline in the United Kingdom comprised largely species 
with high vagility, such as the orange tip, Anthocharis cardamines. 

In somewhat oversimplified terms, vagility is a basic characteristic of all organisms, 
providing individuals with options as burgeoning numbers at source increase competition 
for food supplies. It also improves the chances of meeting and mating with an individual 
from another parent stock and increasing the heterozygosity of the gene base of the 
offspring. (To take this any further would lead to a discussion of the value of sexual 
reproduction.) The ability of individuals on the move to respond to certain environmental 
cues of immediate importance, such as a change in air temperature, land in sight, ’ “food 
in sight,” “mate in sight,’ can over-ride the general stimulus to keep moving. The actual 
mechanisms that both initiate and cease migration are probably hormonal in nature, even 
if the basis is genetic. 

If there is now no return migration to the southeastern United States by resident monarch 
butterflies in Bermuda, it may be that the local cues, e.g., somewhat less fluctuation in 
photoperiod than in New England or southern Canada, or the winter being mild enough 
for adult survival, may serve to suppress the instinct to leave. Perhaps the prevailing 
seasonal winds around Bermuda may make it a one-way journey for all immigrant 
Lepidoptera whatever their migratory indications. Has it been established that all Ber- 
muda monarchs are now non-migratory, or is this an assumption (reasonable enough on 
the face of it) based on monarchs or their immature stages being found there year-round? 
It would be useful to clarify this point. It also would be interesting to release Bermuda 
monarchs in critical localities in North America and see what they would do, for example, 
in the face of a much reduced day-length in central Ontario in early autumn. In a few 
years time, developments in “wildlife technology” will make radio emitters approaching 
the size of a pinhead commercially available for this kind of study where mass alar tagging 
is not appropriate. Prototypes already have been tested. 

I think it unlikely that long distance return migrations commonly evolve from long 
distance one-way dispersals, not just in the case of Bermuda but also on any continental 
mass. It seems to need too many unlikely events to occur in sequence. Initial returns 
probably would be accidental. Development of migration in this way would require not 
only that a few founders somehow find their way back carrying the behavioral trait with 
them, but also that over a number of generations the survival rate of their offspring should 
exceed that extant in the general population. In the first few generations the statistical 
chances of their elimination would seem to be high, unless conditions specifically favorable 
to the migrators developed quite rapidly in the source habitat. 


VOLUME 48, NUMBER 4 391 


More realistically, migrations can generally be considered as a survival mechanism 
evolved, probably during periods of climatic and/or ecological change, as a response to 
gradual increase in segregation or ““de-coupling’’ of optimal zones for feeding and breeding 
which had been more closely adjacent in the earlier history of the population. No near- 
miraculous return flights need be postulated, only slowly widening, short-distance seasonal 
oscillations. Under these circumstances, natural selection for capacities to navigate and 
recognize cues could occur with plenty of opportunity for the advantageous traits to 
accumulate in the population. In the case of the monarch and species with a similar 
pattern of migration, another vital zone presumably became segregated from both optimal 
feeding and breeding regions, i.e., that in which the species could survive the winter in 
the adult stage. Perhaps the monarch already was locked into this type of life cycle, 
because of critical requirements of one or more of the immature stages that could not 
be easily modified subsequently. The fact that we see this kind of pattern in a number 
of the large Holarctic nymphalids suggests that there has been insufficient time for natural 
selection to favor alternatives other than extension of the seasonal movements and adult 
overwintering. If we look for distinctly warmer regimes, when the monarch might have 
been able to overwinter easily at significantly higher latitudes than today, there seem to 
be three main choices, the rather arid, warm Upper Oligocene, part of the Miocene, or 
the post-glacial “Warm Period” of about 6,000 years ago. In theory, 5-6,000 generations 
does not seem an unreasonable time for significant changes in population behaviors to 
occur, although probably far too short a time for significant changes in biologically 
important physiology or morphology. The exceptions might include wing color patterns, 
which often play a role in behavioral responses and recognition and could be subject to 
quite rapid selection. 

My conclusion then, is that the probable answers to the important questions posed by 
Ferguson in his essay in this monograph are as follows: monarchs and many of the other 
butterflies and moths that reach Bermuda became long distance migrators on the mainland 
in response to historical changes in their habitats, much like the migratory songbirds. We 
don't know when this happened, because behavior does not leave a fossil record. Their 
original ranges may have been much the same as today latitudinally, but with the critical 
difference that the adults could overwinter through much of the range instead of only 
at the southern extremity. The overwintering stage of the life cycle may have been the 
most critical factor in determining which options were open to selection when conditions 
changed. 

It would be interesting to make some experimental comparisons between the critical 
factors that appear to operate in such migrations in North America with those in wet- 
dry season and low-high altitude migrations by Lepidoptera, for example in east Africa 
and India. 

In the case of migrations to Bermuda, the driving forces may be quite incidental; there 
are some species which have relatively great vagility over land. When these are swept 
offshore into air masses with a lower level jet stream, they can remain aloft for long 
periods using the same adaptations that serve when they are over the continent. Because 
the basic dispersal instinct can be over-ridden by immediate stimuli related to basic 
survival, they will surely attempt to descend to feed and rest if an oceanic island is located. 
Prevailing seasonal winds perhaps prevent them from ever returning to southeastern 
regions of the United States, but it would be exceedingly difficult to determine this 
conclusively. Additionally, because they have arrived somewhere with resources, the 
migratory instinct may be suppressed by something as simple as the act of feeding or 
egg-laying. We have only limited information concerning which hormonal or other 
physiological releasers and suppressors may be involved. It might be instructive to put 
aerial nets on ships passing east and north of Bermuda to see what proportion of individuals 
lift off to fly further. Even rather small species of Lepidoptera, such as Nomophila spp. 
have been recorded far out to sea. 

Ferguson's suggestion, that the current high population levels of species such as Spo- 
doptera spp. and Pseudaletia unipuncta probably result from the relatively recent massive 
vegetation changes in North America, seems very reasonable. Despite pesticide use, our 
techniques of crop monoculture assist population growth of some of these insects, and 


392 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


almost guarantee periodic explosive outbreaks. As these huge populations reduce local 
resources, dispersal is strongly favored. Even in moderately r-selected organisms such as 
cutworm moths (cf. herring, with eggs in the tens of thousands), traits for dispersal are 
probably so universal that further intensive selection may not be necessary. The long- 
term survival problem for these moths, however, is not over when they reach new food 
sources on a small, isolated island group such as Bermuda. The century or so of change 
in North American vegetation is clearly not enough time for selection to work in favor 
of significant decrease in individual fecundity, or the kinds of specialized behaviors evolved 
in Heliconius species in resource-limited tropical habitats (Gilbert 1983). The sedentary 
island populations of recent origin are still going to be prone to periodic, chaotic population 
outbreaks and subsequent crashes at times when sources of natural mortality diminish, 
simply because of their potential fecundity. 

This does raise interesting possibilities for other types of research using island Lepi- 
doptera, in addition to the kinds of studies made by Ferguson on migration. Someone 
might like to examine other aspects of the situation. For example, by looking at related 
species and genera on islands with biotas of different and preferably known ages, could 
we test the hypothesis that endemic island Lepidoptera may have evolved lower fecundity 
rates over time than comparable migrant species, becoming more adapted to the limi- 
tations of their food plant and nectaring resources? Alternatively, do the island immigrant 
populations begin to show more diversity in food plant selection than their continental 
counterparts? 

I thank Boyce Drummond for encouraging me to submit this essay, and two anonymous 
reviewers for helpful comments, criticisms, and suggestions. Some of the costs were 
supported by NSERC Grant OPG0005863. 


LITERATURE CITED 


BAKER, R. R. 1984. The dilemma: When and how to go or stay, pp. 279-296. In Vane- 
Wright, R.I. & P. R. Ackery (eds.), The biology of butterflies. Symp. Roy. Entomol. 
Soc. London, Academic Press, London. 

DEMPSTER, J. P. 1989. Insect introductions: Natural dispersal and population persistence 
in insects. The Entomologist 108:5-13. 

DENNIS, R. L. H. 1976. The British butterflies. Their origin and establishment. E.W. 
Classey Ltd., Faringdon, England. 318 pp. 

Douc.Las, M. M. 1986. The lives of butterflies. University of Michigan Press, Ann Arbor, 
i-xv + 24] pp. ; 

EHRLICH, P. R. 1984. The structure and dynamics of butterfly populations. pp. 25-40. 
In Vane-Wright, R. I. & P. R. Ackery, P.R. (eds), The biology of butterflies. Symp. 
Roy. Entmol. Soc. London, Academic Press, London. 

FERGUSON, D. C. 1991. An essay on the long-range dispersal and biogeography of 
Lepidoptera, with special reference to the Lepidoptera of Bermuda, pp. 67-77. In 
Ferguson D. C., D. J. Hilburn & B. Wright, The Lepidoptera of Bermuda: Their 
food plants, biogeography, and means of dispersal. Mem. Entomol. Soc. Canad. 158. 
105 pp. 

FERGUSON, D. C., D. J. HILBURN & B. WRIGHT. 1991. The Lepidoptera of Bermuda: 
Their food plants, biogeography, and means of dispersal. Mem. Entomol. Soc. Canad. 
158. 105 pp. 

GASKIN, D. E. 1993. Book review of Ferguson, Hilburn & Wright (1991). J. Lepid. Soc. 
47:165-167. 

GILBERT, L. E. 1983. Coevolution and mimicry, pp. 263-281. In Futuyma, D. J. & M. 
Slatkin (eds.), Coevolution. Sinauer Associates Inc., Sunderland, MA. 

KuUDRNA, O. 1986. Aspects of the conservation of butterflies in Europe. Butterflies of 
Europe, Vol. 8, AULA-Verlag, Wiesbaden. 323 pp. 

PULLIN, A. S., I. F.G. MCLEAN & M. R. WEBB. 1993. The large copper butterfly: British 
and European perspectives, pg. 10. In Abstracts from “Ecology and conservation of 
butterflies,” an international symposium organized by Butterfly Conservation, 10-12 
September 1993, Keele University, England. 28 pp. 


VOLUME 48, NUMBER 4 393 


RILEY, A. M. & D. K. RILEY. 1992. Southerly movements of the red admiral butterfly, 
Vanessa atalanta L. in 1991. Entomol. Rec. 104:82-83. 

ScoTT, J. A. 1986. The butterflies of North America. A natural history and field guide. 
Stanford Univ. Press, California. 583 pp. 

URQUHART, F. A. & N. R. URQUHART. 1978. Autumnal migration routes of the eastern 
population of the monarch butterfly (Danaus p. plexippus) in North America to the 
over-wintering site in the Neovolcanic Plateau of Mexico. Canad. J. Zool. 56:1754- 
1764. 


DAVID E. GASKIN, Department of Zoology, College of Biological Science, University 
of Guelph, Guelph, Ontario NIG 2W1, Canada. 


Received for publication 25 June 1998; revised and accepted 5 April 1994. 


Journal of the Lepidopterists’ Society 
48(4), 1994, 398-394 


CORRECTIONS TO “CHECK LIST OF THE OLD WORLD 
EPIPASCHIINAE AND THE RELATED NEW WORLD GENERA 
MACALLA AND EPIPASCHIA (PYRALIDAE)” 


Additional key words: Lepidogma latifasciata, Locastra, Stericta kiiensis, Teliphasa 
dibelana, Trichotophysa. 


The following are corrections to a list of Old World Epipaschiinae (Solis 1992) discovered 
after publication. The corrections are listed in alphabetical order as in the original work. 
Included are several corrections pertaining to the Japanese fauna identified by Inoue 
(1993). In addition to the corrections listed below, the valid name for the host plant 
Eugenia jambolana Lamarck (Solis 1992:281) should be Syzgium cumini (Linn.) (G. 
Robinson pers. comm.). 


Coenodomus hampsoni West, 1931: misspelled locality name, should be Palali, not Palili. 

Epilepia dentatum (Matsumara and Shibuya, 1927): country should be Japan, not For- 
mosa. 

Lepidogma latifasciata (Wileman, 1911): it is a new combination. 

Lista plinthochroa West, 1931: misspelled locality name, should be Palali, not Palili. 

Locastra maimonalis (Walker, [1859] 1859): years are [1859] 1858, and it is a junior 
synonym of L. crassipennis (Walker). 

Locastra crassipennis (Walker, 1857): senior synonym of L. maimonalis. 

Neopaschia nigromarginata Viette, 19538: locality should be Madagascar Est: Fianarant- 
soa, not Madagascar, Saigon. 

Odontopaschia economia Turner, 1913: misspelled species name, should be ecnomia. 

Orthaga durranti West, 1931: misspelled locality, should be Philippine Islands, not Phil- 
ippines Islands. 

Salma nyctizonalis (Hampson, 1916a): misspelled locality, should be Philippine Islands, 
not Philippines. 

Shoutedenidea Ghesquiére, 1942: misspelled genus name, should be Schoutendenidea. 

Stericta angulosa de Joannis, 1929: misspelled locality, should be Vietnam, not Vitenam. 

Stericta angulosa de Joannis, 1929: misspelled locality, should be Vietnam, not Vitenam. 

Stericta kiiensis (Marumo, 1920): valid species; is not a junior synonym of Stericta 
rufescens (Marumo, 1920), should be placed after Stericta indistincta. 

Termioptycha elegans (Butler, 1881): species belongs in Teliphasa, should be placed after 
Teliphasa dibelana. 


394 JOURNAL OF THE LEPIDOPTERISTS SOCIETY 


Trichotophysa juncundalis (Walker, 1865): misspelled species name, should be jucun- 
dalis, and correct year should be 1866. 
Trichotophysa yamatomis (Strand, 1919): country is Japan, not Taiwan. 
Incertae sedis zophoptera (Ghesquierer, 1942): misspelled author name, should be Ghes- 
quiére. 
LITERATURE CITED 


INOUE, H. 1993. Review of “Check list of the Old World Epipaschiinae and the related 
New World genera Macalla and Epipaschia (Pyralidae).” Japan Heterocerists’ J. 174: 
431. 


SoLis, M. A. 1992. Check list of the Old World Epipaschiinae and the related New 
World genera Macalla and Epipaschia (Pyralidae). J. Lepid. Soc. 46:280-297. 


M. ALMA SOLIS, Systematic Entomology Laboratory, USDA, % National Museum of 
Natural History, Washington, D.C. 20560, USA. 


Received and accepted for publication 25 April 1994. 


CORRECTIONS TO VOLUMES 47 AND 48 


The caption for the cover illustration of 47(2) should have read: “A 
silverspot butterfly (Speyeria sp.) resting beneath the leaf of a fern. 
Original drawing by Martie Clemons from a photograph ee David 
Liebman.” 


The correct order of authorship for the article “Clinal variation in 
Hesperia leonardus (Hesperiidae) in the Loess Hills of the Missouri 
River Valley,” which appeared in 47(4):291-302, should be Stephen M. 
Spomer, Timothy T. Orwig, Leon G. Higley, Gerald L. Selby, and 
Linda J. Young. 

In the article by Astrid Caldas, “Biology of Anaea ryphea (Nymphal- 
idae) in Campinas, Brazil,’ which appeared in 48(3):248-257, the sub- 
specific name of Anaea troglodyta borinquenalis was misspelled as 
“borquenalis’ on page 255. 


The Editor apologizes for these mistakes and any others that have not 
been brought to my attention. 


Journal of the Lepidopterists’ Society 
48(4), 1994, 395-399 


INDEX FOR VOLUME 48 


(New names in boldface) 


acesias, Heliothis, 108 
acrocnema, Boloria, 173 
Acrolepia, 46 
Acrolepiidae, 46 
Acrolepiopsis 

californica, 46 

liliivora, 46 
admirals, 190, 199 
Aglais urticae, cover illustration 48(3) 
agnesaé, Quasimellana, 273 
Agraulis vanillae, 373 
Alliaria petiolata, 168, 171 
amicus, Quasimellana, 274 
ammonia, 205 
Amphipyrinae, 121 
Anaea ryphea, 248 
Anatrytone 

barbara, 273 

flavens, 273 

gladolis, 273 

logan, 278 

mazai, 273 

mella, 273 

perfida, 273 

potosiensis, 273 

sarah, 273 
Andaman Islands, 111 
Andean, 180 
andersoni, Quasimellana, 274 
angra, Quasimellana, 274 
antipazina, Quasimellana, 273 
ants, 381 
Apamea 

ophiogramma, 121 

unanimis, 121 
A practical guide to butterflies and moths 

in southern Africa (book review), 
82 

archippus, Limenitis, 190, 199 
archippus floridensis, Limenitis, 190, 199 
argos, Atrytone, 273 
Aristolochiaceae, 111 
aristolochiae, Pachliopta, 111 
Artemisia dracunculus, 68 
arthemis, Limenitis, 190, 199 
arthemis astyanax, Limenitis, 190, 199 
Atrytone argos, 273 
aurifusa, Josia, 361 
aurora, Quasimellana, 273 
australis, Heliothis, 108 
avian predation, 148 


balsa freemani, Quasimellana, 273 
baracoa, Polites, 37 


barbara, Anatrytone, 273 

Barbehenn, R. V., 386 

Basilarchia, 190, 199 

Bauza, J. A. T., 266 

behavior, 157 

Bermuda, 388 

bilateral size asymmetry, 264 

biological indicators, 173 

biology, 248, 386 

Boloria acrocnema, 173 

bombycoides, Lapara, 51 

Borkin, S. S., 267 

brachypterous, 8 

Brassicaceae, 168, 171 

Brazil, 248 

Britten, H. B., 173 

Brou, V. A., 51 

Brower, A. V. Z., 166 

Burns, J. M., 24, 273 

butterflies, 228 

Butterflies of Tanzania (book review), 80 

butterfly conservation, 111 

Butterfly gardening in the south: Cultivat- 
ing plants that attract butter- 
flies (book review), 76 


Caldas, A., 248 

Calhoun, J. V., 58 
California, 8, 148 
californica, Acrolepiopsis, 46 
Calvert, W. H., 157 
Campinas, Brazil, 248 
Camponotus floridanus, 381 
Canada, 85 

cardenolides, 148 
Caribbean, 267 

carus, Polites, 37 

chamelai, Oidaematophorus, 71 
character reduction, 24 
character loss, 24 
charithonia, Heliconius, 166 
charitonia, Heliconius, 166 
checklist, 228, 393 

chemical defense, 381 
Chemsak, J. A., 71 

Chew, F. S., 168 

China, 128 

cladistics, 129 

clavus, Mellana, 274 
clytemnestra, Hypna, 248 
collecting equipment, 205 
Colorado, 178 

coniferarum, Lapara, 51 
conservation, 111, 171, 178 


396 JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


coon, Pachliopta, 111 food plants, 338, 386 

Cordia, 72 fujisanus, Sibataniozephyrus, 129 
Costa Rica, 72 fuscus, Diaphanos, 180 

county records, 1 

Courant, A. V., 168 Gaedike, R., 46 

Covell, C. V., 199 gala, Mellana, 274 

Croton, 72, 248 Gaskin, D. E., 388 

cryptic species, 106 generic limits, 24 

curvignathos, Diaphanos, 180 genetics, 264 


cybele, Speyeria, cover illustration 48(4) genitalia, 18, 25, 47, 53, 69, 107, 132, 273 
geographic distribution, 273 


Dalceridae, 381 Geometridae, 8 
Dalcerides ingenita, 381 Gibson, L. D., 69 
Danainae, 148, 157 giulianiata, Tescalsia, 12 
Danaus plexippus, 148, 157 gladolis, Anatrytone, 273 
Diaphanos gopala, Josia, 357 
curvignathos, 180 grasses, 386 
fuscus, 180 Great Plains, 228 
huberi, 180 
diet breadth, 168 habitat suitability, 173 
Digitivalva, 46 halicarnie, Lapara, 51 
Dioptinae, 338 Hardwick, D. F., 106 
discinota, Thirmida, 345 hawkmoths, 51 
Dismorphia spio, 266 hector, Pachliopta, 111 
Dismorphiinae, 266 Heliconiinae, 166 
Disporum hookeri, 49 Heliconius 
distribution, 1, 24, 72, 180, 228, 273 charithonia, 166 
diurnal moth, 71 charitonia, 166 
double-mount, 205 Heliothentinae, 106 
Heliothis 
early stages, 252, 381 acesias, 108 
Eisner, T., 381 australis, 108 
endangered species, 173 phloxiphaga, 106 
endemic species, 180 helva, Mellana, 274 
endemism, 51 Hesperiidae, 24, 273, 386: 
Etheridge, M., cover illustration 48(4) Hesperioidea, 228 
Epipaschia, 393 Hispanola, 267 
Epipaschiinae, 393 Holbrook, A. E., 168 
Epstein, M. C., 381 host expansion, 171 
Eucosma host plant, 68, 48, 168, 171, 338, 386 
ustulatana, 69 Hsu, Y.-F., 128 
uta, 69 huberi, Diaphanos, 180 
wandana, 69 hybridization, 190, 199 
eulogius, Quasimellana, 273 Hymenoptera, 266 
Euphorbiaceae, 248 Hypna clytemnestra, 248 
evolution, 388 
external morphology, 258 immatures, 338 
imperfida, Quasimellana, 274 
Fagus-feeding, 128 inconspicua, Quasimellana, 274 
Fairyland, 59 India, 111 
Ferguson, D. C., 8 Indian River, 58 
fieldi, Quasimellana, 274 indra, Papilio, 68 
Filbert, M. C., cover illustration 48(2) ingenita, Dalcerides, 381 
flavens, Anatrytone, 273 integumental coating, 381 
flightless, 8 introductions, 121, 168, 171 


flight period, 231 
Florida, 58, 373 Japan, 128 


VOLUME 48, NUMBER 4 


Josia 
aurifusa, 361 
gopala, 357 
radians, 353 
turgida, 363 
Josiini, 338 
Junonia coenia, cover illustration 48(2) 
jurtina, Maniola, 264 


Kentucky, 69 

Key to the insects of the European part of 
the USSR, partIV (Lepidoptera) 
(book review), 68 

kiiensis, Stericta, 393 

kuafui, Sibataniozephyrus, 143 


Lafontaine, J. D., 121 
Landry, B., 205 
Landry, J.-F., 205 
Lapara 
bombycoides, 51 
coniferarum, 51 
halicarnie, 51 
phaeobrachycerous, 51 
pinea, 51 
Larentiinae, 8 
larvae, 248, 258, 338, 381 
larval morphology, 258, 338 
Lepidogma latifasciata, 393 
life history, 19, 58, 111, 248, 248, 338 
Lilium washingtonianum, 48 
liliivora, Acrolepiopsis, 46 
Limenitis 
archippus, 190, 199. 
archippus floridensis, 190, 199 
arthemis, 190, 199 
arthemis astyanax, 190, 199 
Lin, M.-Y., 128 
Locastra, 398 
logan, Anatrytone, 273 
log-series model, 85 
Louisiana, 51 
Lycaenidae, 128 


Macalla, 393 
machaon bairdii, Papilio, 68 
Maniola 

jurtina, 264 
Marrone, G. M., 228 
Maudsley, J. R., 190 
mazai, Anatrytone, 273 
McKown, S., 82 
mella, Anatrytone, 273 
Mellana, 273 
Memphis, 248 
meridiani, Quasimellana, 274 
mexicana, Quasimellana, 273 


397 


Mexico, 24, 72, 157 
microlepidoptera, 205 
mielkei, Quasimellana, 274 
migration, 373, 388 
migratory cues, 388 
Mikkola, K., 121 

Miller, J. S., 338 

Miller, J. Y., 76 

Miller, L. D., 80 

Miller, W. E., 69 

mimicry, 71, 190 

minata, Tescalsia, 21 
Minet, J., 74 

Mohanraj, P., 111 

monarch, 148, 157 

monica, Quasimellana, 274 
montane forest, 338 
montezuma, Quasimellana, 274 
mosaicism, 264 

mounting technique, 205 
mulleri, Quasimellana, 273 
Musineon tenuifolia, 68 
mustard, 168, 171 

myron, Quasimellana, 274 
mystic, Polites, 24 


napi, Pieris, 169 

nayana, Quasimellana, 273 

Nearctic, 69 

Nebraska, 68 

nectar source, 173 

Nevada, 8 

New World, 273, 393 

Nicobar Islands, 111 

nicomedes, Quasimellana, 274 

Nielsen, M. C., 79 

Noctuidae, 106, 121 

noka, Quasimellana, 273 

nomenclature, 166 

North America, 121 

Notodontidae, 338 

Nymphalidae, 148, 157, 166, 173, 180, 190, 
199, 248, 259, 264 


oaxaca, Quasimellana, 273 
obituary, 267 
Oidaematophorus 

chamelai, 71 
Old World, 393 
Olethreutinae, 69 
onychophorans, 381 
ophiogramma, Apamea, 121 
origenes, Polites, 24 
Ostern, H. W., cover illustration 48(3) 
Otero, L. D., 338 
otho, Wallengrenia, 274 
overwintering, 148, 157 


398 


Pachliopta 
aristolochiae, 111 
coon, 111 
hector, 111 
rhodifer, 111 
Palearctic, 121 
pandora, Quasimellana, 274 
Papilio 
indra, 68 
machaon bairdii, 68 


polyxenes, cover illustration 48(1), 68 


zelicaon, 258 

zelicaon nitra, 68 
Papilionidae, 68, 111 
Papilionoidea, 228 
paramo, 180 
parasitoids, 266 
partial-night sampling, 85 
Passiflora, 338 
pazina, Quasimellana, 273 
perfida, Anatrytone, 273 
petiolata, Alliaria, 168, 171 
phaeobrachycerous, Lapara, 51 
Phalaris, 121 
Phoebis sennae, 373 
Phragmites, 121 
phyloxiphaga, Heliothis, 106 
Pieridae, 168, 171, 266 
Pieris 

napi, 169 

virginiensis, 168, 171 
pinea, Lapara, 51 
Platt, A. P., 190 
plexippus, Danaus,148, 157 
plume moth, 71 
Poaceae, 386 
Poanes melane, 386 
Poanopsis, 24 
Polites 

baracoa, 37 

carus, 37 

mystic, 24 

origenes, 24 

pupillus, 25 

puxillius, 25 

rhesus, 36 

subreticulata, 37 

themistocles, 25 

vibex, 25 


polyxenes, Papilio, cover illustration 48(1), 


Porter, A., 171 

potosiensis, Anatrytone, 273 
Powell, J. A., 8, 71 
predation, 148, 157 
Presidential Address, 1 
Profile, 58 


JOURNAL OF THE LEPIDOPTERISTS’ SOCIETY 


Pronophilini, 180 
protean behavior, 157 
Pterophoridae, 71 
Puerto Rico, 266 
pupillus, Polites, 25 
puxillius, Polites, 25 
Pyralidae, 393 


Quasimellana 
agnesae, 273 
amicus, 274 
andersoni, 274 
angra, 274 
antipazina, 273 
aurora, 273 
balsa freemani, 273 
eulogius, 273 
fieldi, 274 
imperfida, 274 
inconspicua, 274 
meridiani, 274 
mexicana, 273 
mielkei, 274 
monica, 274 
montezuma, 274 
mulleri, 273 
myron, 274 
nayana, 273 
nicomedes, 274 
noka, 273 
oaxaca, 273 
pandora, 274 
pazina, 273 
sethos, 274 
siblinga, 273 
sista, 273 
tamana, 273 
tecla, 273 
verba, 274 

Quercusia, 128 


radians, Josia, 358 
Reiser, J. M., 68 

rhesus, Polites, 24 
Rhizedra lutosa, 121 
rhodifer, Pachliopta, 111 
Riley, L., 173 

riparian moths, 121 
rivoula, Mellana, 274 
“rubidus,’’ 190 
rufous-sided towhee, 148 
ryphea, Anaea, 248 


Sakai, W. H., 148 
sampling strategies, 85 
sarah, Anatrytone, 273 
Satyrinae, 180, 264 


VOLUME 48, NUMBER 4 


Schwartz, A., 267 
selection, 190 
sethos, Quasimellana, 274 
setting and mounting, 205 
sex determination, 258 
sexing larvae, 258 
sex ratios, 148 
Sibataniozephyrus 

fujisanus, 129 

kuafui, 143 
siblinga, Quasimellana, 273 
sista, Quasimellana, 273 
sister species, 273 
skippers, 24, 228, 386 
Smedley, S. R., 381 
Solis, M. A., 393 
South Dakota, 228 
speciation, 24 
species-abundance distribution, 85 
species diversity, 85 
species group, 273 
spelling, 166 
Speyeria cybele, cover 48(4) 
Sphingidae, 51 
Spomer, S. M., 68 
spreading box, 205 
staging, 205 
Stanford, R. E., 1 
startle response, 157 
Stericta kiiensis, 393 
subreticulata, Polites, 37 
systematics, 128, 273 
Szafranski, P., 264 


tagging, 148 
Taiwan, 128 
tamana, Quasimellana, 273 
taxonomy, 106, 166 
techniques, 205, 258 
tecla, Quasimellana, 273 
Teliphasa, 393 
Tescalsia 
giulianiata, 12 
minata, 21 
The butterflies of Kenya and their natural 
history (book review), 80 
Theclinae, 128 
Theclini, 128 
themistocles, Polites, 25 
The Ontario butterfly atlas (book review), 
79 
Thirmida 
discinota, 345 


399 


Thomas, A. W., 85 
Thomas, G. M., 85 
Torres Bauza, J. A., 266 
Tortricidae, 69 

trap, 373 

trapping, 9, 85, 373 
Trichogramma, 266 
Trichospilus, 266 
Trichotophysa, 398 
Troidini, 111 
turgida, Josia, 363 
tympana, 8 

type locality, 58 


Uncompahgre fritillary, 173 
Underwood, D. L. A., 258 

unanimis, Apamea, 121 

United States, 190, 199 

urticae, Aglais, cover illustration 48(3) 
ustulatana, Eucosma, 69 

uta, Eucosma, 69 


Van der Reijden, E. D., 168 
vanillae, Agraulis, 373 
Veenakumari, K., 111 
Venezuela, 180, 338 

verba, Quasimellana, 274 
vials, 205 

vibex, Polites, 25 

Viloria P., A. L., 180 
virginiensis, Pieris, 168, 171 
voltinism, 51 


Wallengrenia 
otho, 274 
Walker, T. J., 373 
wandana, Eucosma, 69 
Whitesell, J. J., 373 
Wild, E. R., cover illustration 48(1) 
winter moths, 8 
Wittfeld, Annie, 58 
Wittfeld, William, 58 


Yponomeutoidea, 46 
Yvretta, 24 


zelicaon, Papilio, 258 
zelicaon nitra, Papilio, 68 


30-minute samples, 85 


Date of Issue (Vol. 48, No. 4): 19 December 1994 


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EDITORIAL STAFF OF THE JOURNAL 
JOHN W. Brown, Editor 
Entomology Department 
San Diego Natural History Museum 
P.O. Box 1390 
San Diego, California 92112, U.S.A. 


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CHRISTER WIKLUND (Sweden) 


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CONTENTS 


GENITALIA AT THE GENERIC LEVEL: ATRYTONE RESTRICTED, ANATRY- 
TONE RESURRECTED, NEW GENUS QUASIMELLANA—AND YES! WE 
HAVE NO MELLANAS (HESPERIUDAE) John M. Burns -——---—-—— De 


IMMATURE STAGES OF VENEZUELAN DIOPTINAE (NOTODONTIDAE) IN 
JOSIA AND THIRMIDA James S. Miller and L. Daniel Otero __ 338 


A SIMPLE PORTABLE TRAP FOR MIGRATING BUTTERFLIES Thomas E 
Walker and James J, Whitesell.) as) hs 


GENERAL NOTES | 


Sticky integumental coating of a dalcerid caterpillar: A deterrent to ants Mare - 


E. Epstein, Scott’ R, Smedley and Thomas Hisner 2 |) st 381 

Host plants of Poanes melane (Hesperiidae) Raymond V. Barbehenn ___. 386 
Comments on the nature and origins of migrations of Lepidoptera to Bermuda 

David EMGaskinar | ye a AS SS Ss Ee 388 
Corrections to “Check list of the Old World Epipaschiinae and the related New 

World genera Macalla and Epipaschia (Pyralidae)” M. Alma Solis ---------- 393 

CORRECTIONS TO VOLUMES 4 7:AND 48 (= Ee 394 

INDEX#OR VOU NG)4 0) ee 395 


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CAIN APS WAU conga aie aS AsBeAye aye 
YOAR SS OR Re ve 


CONE SGA Nay eg, 


TRL PW pay 


Pye 


we Atv 


diel